Fungi and Food Spoilage, 3rd ed

Fungi and Food Spoilage

“This page left intentionally blank.”

John I. Pitt

l

Ailsa D. Hocking

Fungi and Food Spoilage

13

John I. Pitt Honorary Research Fellow CSIRO Food and Nutritional Sciences North Ryde, NSW 2113 Australia [email protected]

Ailsa D. Hocking Honorary Research Fellow CSIRO Food and Nutritional Sciences North Ryde, NSW 2113 Australia [email protected]

ISBN 978-0-387-92206-5 e-ISBN 978-0-387-92207-2 DOI 10.1007/978-0-387-92207-2 Springer Dordrecht Heidelberg London New York Library of Congress Control Number: 2009920217 # Springer ScienceþBusiness Media, LLC 2009 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Springer ScienceþBusiness Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper Springer is part of Springer ScienceþBusiness Media (www.springer.com)

Preface to the Third Edition

In contrast to the second edition, the third edition of ‘‘Fungi and Food Spoilage’’ is evolutionary rather than revolutionary. The second edition was intended to cover almost all of the species likely to be encountered in mainstream food supplies, and only a few additional species have been included in this new edition. The third edition represents primarily an updating – of taxonomy, physiology, mycotoxin production and ecology. Changes in taxonomy reflect the impact that molecular methods have had on our understanding of classification but, it must be said, have not radically altered the overall picture. The improvements in the understanding of the physiology of food spoilage fungi have been relatively small, reflecting perhaps the lack of emphasis on physiology in modern microbiological science. Much remains to be understood about the specificity of particular fungi for particular substrates, of the influence of water activity on the growth of many of the species treated, and even on such basic parameters as cardinal temperatures for growth and the influence of pH and preservatives. Since 1997, a great deal has been learnt about the specificity of mycotoxin production and in which commodities and products-specific mycotoxins are likely to occur. Changes in our understanding of the ecology of the included species are also in most cases evolutionary. A great number of papers have been published on the ecology of foodborne fungi in the past few years, but with few exceptions the basic ecology of the included species remains. Recent changes in our understanding of foodborne fungi include the realisation that Aspergillus carbonarius is a major source of ochratoxin A in the world food supply, that A. westerdijkiae and not A. ochraceus is the other common Aspergillus species making this toxin and that these species are responsible for ochratoxin A in foods outside the cool temperate regions, where Penicillium verrucosum is the important species. In recent years a number of new species have been found to be capable of producing aflatoxin, but the fact remains that most aflatoxin in the global food supply is produced by A. flavus and A. parasiticus. The taxonomy of Fusarium species is still undergoing major revision. However, the renaming of Fusarium moniliforme as F. verticillioides is the only change of importance here. Recent publications have improved our understanding of species – mycotoxin relationships within Fusarium.

v

vi

Among the colleagues who helped us to prepare this edition, we wish to particularly thank Dr Anne-Laure Markovina, now of the University of Sydney, who assisted in literature searches and some cultural and photographic work, and Mr N.J. Charley who has continued his excellent work of curating the FRR culture collection, on which so much of the descriptive work in this book is based.

Preface to the Third Edition

Preface to the First Edition

This book is designed as a laboratory guide for the food microbiologist to assist in the isolation and identification of common foodborne fungi. We emphasise the fungi which cause food spoilage, but also devote space to the fungi commonly encountered in foods at harvest, and in the food factory. As far as possible, we have kept the text simple, although the need for clarity in the descriptions has necessitated the use of some specialised mycological terms. The identification keys have been designed for use by microbiologist with little or no prior knowledge of mycology. For identification to genus level, they are based primarily on the cultural and physiological characteristics of fungi grown under a standard set of conditions. The microscopic features of the various fungi become more important when identifying isolates at the species level. Nearly all of the species treated have been illustrated with colony photographs, together with photomicrographs or line drawings. The photomicrographs were taken using a Zeiss WL microscope fitted with Nomarski interference contrast optics. We are indebted to Mr W. Rushton and Ms L. Burton, who printed the many hundreds of photographs used to make up the figures in this book. We also wish to express our appreciation to Dr D.L. Hawksworth, Dr A.H.S. Onions and Dr B.C. Sutton of the Commonwealth Mycological Institute, Kew, Surrey, UK, Professor P.E. Nelson and the staff of the Fusarium Research Center, University of Pennsylvania, USA and Dr L.W. Burgess of the University of Sydney, who generously provided facilities, cultures and advice on some of the genera studied.

vii

“This page left intentionally blank.”

Preface to the Second Edition

In planning for the second edition of ‘‘Fungi and Food Spoilage’’, we decided that the book would benefit from a larger format, which would permit improved illustrations, and from some expansion of the text, in both numbers of species treated and overall scope. These aims have been realised. The Crown Quarto size has allowed us to include substantially larger, clearer illustrations. Many new photographs and photomicrographs have been added, the latter taken using a Zeiss Axioscop microscope fitted with Nomarski differential interference contrast optics. We have taken the opportunity to include more than 40 additional species descriptions, to add a new section on mycotoxin production for each species and to update and upgrade all of the text. Since the first edition, changes in the climate for stabilising fungal nomenclature have resulted in development of a list of ‘‘Names in Current Use’’ for some important genera, including Aspergillus and Penicillium. Names of species used in the second edition are taken from that list, which was given special status by the International Botanical Congress, Tokyo, 1994. Names used in this edition have priority over any other names for a particular species. Publication of a list of ‘‘Authors of Fungal Names’’ (P.M. Kirk and A.E. Ansell, Index of Fungi, Supplement: 1–95, 1992) has also stabilised names of authorities for all fungal species. Abbreviations of authors’ names used in this edition conform to those recommended by Kirk and Ansell. Some progress in standardisation of methods and media has also been made, primarily through the efforts of the International Commission on Food Mycology. The first edition included some 400 references. When we began revisionary work, we felt that the number of references in the area of food mycology had probably doubled or increased by perhaps 150% during the intervening years. In fact, this second edition includes over 1900 references, almost a five-fold increase over the 1985 edition! This provides a clear indication that interest in, and study of, food mycology has greatly increased in recent years. Modern referencing systems have enabled us to expand information from tropical sources, especially in Asia and Africa, but we are conscious of the fact that treatment of fungi found in foods on a worldwide basis remains rather incomplete. We gratefully acknowledge support and assistance from colleagues who have contributed to this new edition. Ms J.C. Eyles formatted and printed the camera

ix

x

ready copy, Ms C. Heenan collated, arranged and formatted the illustrations and Mr N.J. Charley looked after the culture collection, culture growth and colony photography. Without this level of support, the book would not have been completed.

Preface to the Second Edition

Contents 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1

2

The Ecology of Fungal Food Spoilage . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Water Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Hydrogen Ion Concentration . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Temperature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Gas Tension . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Consistency. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Nutrient Status . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Specific Solute Effects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.8 Preservatives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 Conclusions: Food Preservation . . . . . . . . . . . . . . . . . . . . . . . .

3 3 4 5 7 8 8 8 9 9

3

Naming and Classifying Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Taxonomy and Nomenclature: Biosystematics . . . . . . . . . . . . . 3.2 Hierarchical Naming . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Zygomycotina. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Ascomycotina . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5 Basidiomycotina . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.6 The Ascomycete – Conidial Fungus Connection . . . . . . . . . . . . 3.7 Dual Nomenclature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.8 Practical Classification of Fungi . . . . . . . . . . . . . . . . . . . . . . . .

11 11 12 12 13 15 15 15 16

4

Methods for Isolation, Enumeration and Identification . . . . . . . . . . . . 4.1 Sampling. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Enumeration Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.1 Direct Plating. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.2 Dilution Plating . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.3 Incubation Conditions. . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Sampling Surfaces . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Air Sampling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5 Isolation Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.1 Yeasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.2 Moulds. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.3 Short Term Storage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6 Choosing a Suitable Medium . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6.1 General Purpose Enumeration Media. . . . . . . . . . . . . . . 4.6.2 Selective Isolation Media. . . . . . . . . . . . . . . . . . . . . . . . . 4.6.3 Techniques for Yeasts . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6.4 Techniques for Heat-Resistant Fungi . . . . . . . . . . . . . . . 4.6.5 Other Plating Techniques . . . . . . . . . . . . . . . . . . . . . . . . 4.7 Estimation of Fungal Biomass . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7.1 Chitin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7.2 Ergosterol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7.3 Impedimetry and Conductimetry . . . . . . . . . . . . . . . . . .

19 19 19 20 21 22 22 23 23 23 24 24 25 26 27 30 32 33 34 34 35 37 xi

xii

5

Contents

4.7.4 Adenosine Triphosphate (ATP) . . . . . . . . . . . . . . . . . 4.7.5 Fungal Volatiles. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7.6 Immunological Techniques . . . . . . . . . . . . . . . . . . . . . 4.7.7 Molecular Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.8 Identification Media and Methods. . . . . . . . . . . . . . . . . . . . . . 4.8.1 Standard Methodology . . . . . . . . . . . . . . . . . . . . . . . . 4.8.2 Plating Regimen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.8.3 Inoculation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.8.4 Additional Media and Methods . . . . . . . . . . . . . . . . . 4.8.5 Identification of Fusarium Species. . . . . . . . . . . . . . . . 4.8.6 Yeasts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.9 Examination of Cultures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.9.1 Colony Diameters . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.9.2 Colony Characters. . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.9.3 Preparation of Wet Mounts for Microscopy. . . . . . . . 4.9.4 Staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.9.5 Microscopes and Microscopy . . . . . . . . . . . . . . . . . . . 4.10 Preservation of Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.10.1 Lyophilisation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.10.2 Other Storage Techniques . . . . . . . . . . . . . . . . . . . . . . 4.11 Housekeeping in the Mycological Laboratory. . . . . . . . . . . . . 4.11.1 Culture Mites. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.11.2 Problem Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.11.3 Pathogens and Laboratory Safety . . . . . . . . . . . . . . . .

37 37 38 40 41 41 41 41 42 43 44 45 45 45 46 46 47 48 48 49 50 50 51 51

Primary Keys and Miscellaneous Fungi . . . . . . . . . . . . . . . . . . . . . . . . 5.1 The General Key . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1.1 Notes on the General Key . . . . . . . . . . . . . . . . . . . . . . 5.2 Miscellaneous Fungi. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3 Genus Acremonium Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.4 Genus Alternaria Nees: Fr.. . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.5 Genus Arthrinium Kunze . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.6 Genus Aureobasidium Viala and G. Boyer . . . . . . . . . . . . . . . . 5.7 Genus Bipolaris Shoemaker . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.8 Genus Botrytis P. Micheli: Fr. . . . . . . . . . . . . . . . . . . . . . . . . . 5.9 Genus Chaetomium Kunze . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.10 Genus Chrysonilia Arx . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.11 Genus Cladosporium Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.12 Genus Colletotrichum Corda . . . . . . . . . . . . . . . . . . . . . . . . . . 5.13 Genus Curvularia Boedijn. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.14 Genus Drechslera S. Ito . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.15 Genus Endomyces Reess . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.16 Genus Epicoccum Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.17 Genus Fusarium Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.18 Genus Geotrichum Link: Fr.. . . . . . . . . . . . . . . . . . . . . . . . . . . 5.19 Genus Hyphopichia Arx and van der Walt. . . . . . . . . . . . . . . . 5.20 Genus Lasiodiplodia Ellis and Everh. . . . . . . . . . . . . . . . . . . . . 5.21 Genus Monascus Tiegh. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.22 Genus Moniliella Stolk and Dakin . . . . . . . . . . . . . . . . . . . . . . 5.23 Genus Nigrospora Zimm. . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

53 54 55 56 58 60 64 65 67 68 70 73 75 81 82 85 86 88 89 122 124 125 127 129 131

Contents

xiii

5.24 5.25 5.26 5.27 5.28 5.29 5.30

Genus Pestalotiopsis Steyaert . . . . . . . . . . . . . . . . . . . . . . . . . . Genus Phoma Sacc. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genus Stemphylium Wallr. . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genus Trichoconiella B.L. Jain. . . . . . . . . . . . . . . . . . . . . . . . . Genus Trichoderma Pers. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genus Trichothecium Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genus Ulocladium Preuss . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

133 134 136 137 139 140 142

6

Zygomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1 Genus Absidia Tiegh. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2 Genus Cunninghamella Matr. . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3 Genus Mucor P. Micheli: Fr. . . . . . . . . . . . . . . . . . . . . . . . . . . 6.4 Genus Rhizomucor (Lucet and Costantin) Vuill. . . . . . . . . . . . 6.5 Genus Rhizopus Ehrenb.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.6 Genus Syncephalastrum J. Schrot. ¨ ...................... 6.7 Genus Thamnidium Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

145 148 149 151 157 158 165 167

7

Penicillium and Related Genera . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1 Genus Byssochlamys Westling . . . . . . . . . . . . . . . . . . . . . . . . . 7.2 Genus Eupenicillium F. Ludw. . . . . . . . . . . . . . . . . . . . . . . . . . 7.3 Genus Geosmithia Pitt . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.4 Genus Paecilomyces Bainier . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.5 Genus Scopulariopsis Bainier . . . . . . . . . . . . . . . . . . . . . . . . . . 7.6 Genus Talaromyces C.R. Benj.. . . . . . . . . . . . . . . . . . . . . . . . . 7.7 Genus Penicillium Link. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.7.1 Penicillium subgenus Aspergilloides Dierckx . . . . . . . . . 7.7.2 Penicillium subgenus Furcatum Pitt. . . . . . . . . . . . . . . . 7.7.3 Penicillium subgenus Penicillium . . . . . . . . . . . . . . . . . . 7.7.4 Penicillium subgenus Biverticillium Dierckx . . . . . . . . .

169 170 175 182 183 187 188 194 196 207 223 263

8

Aspergillus and Related Teleomorphs. . . . . . . . . . . . . . . . . . . . . . . . . . 8.1 Genus Emericella Berk. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2 Genus Eurotium Link: Fr. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.3 Genus Neosartorya Malloch and Cain . . . . . . . . . . . . . . . . . . . 8.4 Genus Aspergillus Fr.: Fr. . . . . . . . . . . . . . . . . . . . . . . . . . . . .

275 279 281 292 295

9

Xerophiles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.1 Genus Basipetospora G.T. Cole and W.B. Kendr. . . . . . . . . . . 9.2 Genus Chrysosporium Corda . . . . . . . . . . . . . . . . . . . . . . . . . . 9.3 Genus Eremascus Eidam . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.4 Genus Polypaecilum G. Sm. . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.5 Genus Wallemia Johan-Olsen. . . . . . . . . . . . . . . . . . . . . . . . . . 9.6 Genus Xeromyces L.R. Fraser . . . . . . . . . . . . . . . . . . . . . . . . .

339 340 342 347 348 350 353

10

Yeasts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

357

11

Fresh and Perishable Foods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.1 Spoilage of Living, Fresh Foods . . . . . . . . . . . . . . . . . . . . . . . 11.2 Fruits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.1 Citrus Fruits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.2 Pome Fruits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.3 Stone Fruits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

383 383 383 384 385 386

xiv

Contents

11.3

11.4 11.5 11.6

12

11.2.4 Tomatoes and other Solanaceous Fruit. . . . . . . . . . . 11.2.5 Melons and other Cucurbits . . . . . . . . . . . . . . . . . . . 11.2.6 Grapes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.7 Berries . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.8 Figs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.9 Tropical Fruit . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Vegetables . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.1 Peas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.2 Beans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.3 Onions and Garlic . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.4 Potatoes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.5 Roots and Tubers . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.6 Yams . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.7 Cassava . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.8 Leafy and other Green Vegetables . . . . . . . . . . . . . . Dairy Foods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Meats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cereals, Nuts and Oilseeds . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.1 Wheat, Barley and Oats. . . . . . . . . . . . . . . . . . . . . . . 11.6.2 Rice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.3 Maize . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.4 Soybeans and Mung Beans . . . . . . . . . . . . . . . . . . . . 11.6.5 Other Beans and Pulses . . . . . . . . . . . . . . . . . . . . . . . 11.6.6 Sunflower Seed . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.7 Sorghum. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.8 Peanuts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.9 Cashews and Brazil Nuts . . . . . . . . . . . . . . . . . . . . . . 11.6.10 Almonds, Hazelnuts, Walnuts and Pecans . . . . . . . . 11.6.11 Pistachios . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.12 Copra . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Spoilage of Stored, Processed and Preserved Foods . . . . . . . . . . . . . . . 12.1 Low Water Activity Foods: Dried Foods . . . . . . . . . . . . . . . . 12.1.1 Cereals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.2 Flour . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.3 Pasta. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.4 Bakery Products. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.5 Maize . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.6 Soybeans, Mung Beans, other Beans and Chickpeas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.7 Nuts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.8 Peanuts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.9 Hazelnuts, Walnuts, Pecans and Almonds . . . . . . . . 12.1.10 Pistachio Nuts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.11 Other Nuts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.12 Coconut . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.13 Spices . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.14 Coffee Beans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.15 Cocoa . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.16 Dried Meat . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

387 387 388 389 389 390 391 391 391 391 392 392 392 392 393 393 394 395 395 396 396 397 398 398 398 398 399 399 399 400 401 401 402 403 403 403 404 405 406 406 407 408 409 409 410 410 411 412

Contents

xv

12.2

Low Water Activity Foods: Concentrated Foods . . . . . . . . . . 12.2.1 Jams . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.2 Dried Fruit. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.3 Fruit Cakes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.4 Confectionery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.5 Fruit Concentrates. . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.6 Honey and Syrups . . . . . . . . . . . . . . . . . . . . . . . . . . . . Low Water Activity Foods: Salt Foods . . . . . . . . . . . . . . . . . . Intermediate Moisture Foods: Processed Meats . . . . . . . . . . . Heat Processed Acid Foods . . . . . . . . . . . . . . . . . . . . . . . . . . . Preserved Foods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cheese . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

412 412 413 414 415 415 416 416 417 418 418 419

Media Appendix . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

423

Glossary. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

427

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

431

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

503

12.3 12.4 12.5 12.6 12.7

Chapter 1

Introduction

From the time when primitive man began to cultivate crops and store food, spoilage fungi have demanded their tithe. Fuzzes, powders and slimes of white or black, green, orange, red and brown have silently invaded – acidifying, fermenting, discolouring and disintegrating, rendering nutritious commodities unpalatable or unsafe. Until recently, fungi have generally been regarded as causing only unaesthetic spoilage of food, despite the fact that Claviceps purpurea was linked to human disease more than 200 years ago, and the acute toxicity of macrofungi has long been known. Japanese scientists recognised the toxic nature of yellow rice 100 years ago, but 70 years elapsed before its fungal cause was confirmed. Alimentary toxic aleukia killed many thousands of people in the USSR in 1944–1947; although fungal toxicity was suspected by 1950, the causal agent, T-2 toxin, was not clearly recognised for another 25 years. Forgacs and Carll (1952), in a prophetic article, warned of the danger from common spoilage fungi, but it was not until 1960, when the famous "Turkey X" disease killed 100,000 turkey poults in Great Britain, and various other disasters followed in rapid succession, that the Western world became aware that common spoilage moulds could produce significant toxins. Since 1960 a seemingly endless stream of toxigenic fungi and potentially toxic compounds has been discovered. On these grounds alone, the statement ‘‘It’s only a mould’’ is no longer acceptable to food microbiologist, health inspector or consumer. The demand for accurate identification and characterisation of food spoilage fungi has become urgent. In the flurry of research into mycotoxins, however, it must not be forgotten that food spoilage as such

remains an enormous problem throughout the world. Figures are difficult to obtain. However, even given a dry climate and advanced technology, losses of food to fungal spoilage in Australia must be in excess of $10,000,000 per annum: losses in humid tropical climates and countries with less highly developed technology remain staggering. An estimate of 5–10% of all food production is not unrealistic. Research into fungal food spoilage and its prevention is clearly an urgent necessity: lacking in spectacular appeal, it is, however, often neglected. A further point, of the highest significance, needs emphasis here. Research on the fungi which cause food spoilage, and the mycotoxins they produce, can only be carried out effectively if based on accurate identification of the microorganisms responsible. Taxonomy and nomenclature (systematics) make up the vital root system of all the trees of biological science. The prevention of fungal food spoilage as an art is old, but as a discipline, young. Drying, the oldest method of food preservation, has been practiced for millennia and is still the most common, effective and cheap technique for preserving food. Only recently have we been able to identify with certainty the fungal species which cause spoilage of dried foods. Prediction of their responses to a given environment, specified by physico-chemical parameters such as water activity, temperature, pH and oxygen tension, even now is often uncertain. Within historic times, newer methods of food preservation have been introduced – salting, curing, canning, refrigeration, freezing, the use of preservatives, irradiation and most recently, high hydrostatic pressure. Freezing excepted, each new technique has selected for one or more fungal species resistant to

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_1, Ó Springer ScienceþBusiness Media, LLC 2009

1

2

the process applied. As examples we can take Polypaecilum pisce on salt fish, Xeromyces bisporus on fruit cake, Cladosporium herbarum on refrigerated meat, Zygosaccharomyces bailii in preserved juices, Z. rouxii in jams and fruit concentrates, Aspergillus flavus on peanuts, Eurotium chevalieri on hazel nuts, Penicillium roqueforti on cheeses, Byssochlamys fulva in acid canned foods . . . the list of quite specific food – fungus associations is extensive. The study of such associations is one of the more important branches of the young discipline, food mycology. This book sets out to document current knowledge on the interaction of foods and fungi, in the context of spoilage and toxicity, not food production or biotechnology. Four aspects are examined. First, ecology: what factors in foods select for particular kinds of fungi? A chapter is devoted to the physical and chemical parameters which influence the growth of fungi in foods. Second, methodology: how do we isolated fungi from foods? What are the best media to use? How do we go about identifying food spoilage fungi? Third, the commodity: what fungi are usually associated with a particular food? Here ecological factors interact to produce a more or less specific habitat. Major classes of foods and their associated spoilage fungi are described. Finally, the fungus: what fungus is that? In a series of chapters, the main food spoilage moulds and yeasts are described and keyed, together with others commonly associated with food but not noted for spoilage. Where possible, further information is

1

Introduction

given on known habitats and sources, physiology, heat resistance, etc., together with a selective bibliography. Accurate information on mycotoxin production is also included. As far as possible, the precise terminology for fungal structures used by the pure mycologist and indeed most necessary for him has been avoided in these chapters. Some concepts and terms are of course essential: these have been introduced as needed and are listed in a glossary. The taxonomic sections of this book are designed to facilitate identification of food spoilage and common food contaminant fungi. A standardised plating regimen is used, originally developed for the identification of Penicillium species (Pitt, 1979b) and extended here to other genera relevant to the food industry. Under this regimen, cultures are incubated for 1 week at 5, 25 and 378C on a single standard medium and at 258C on two others. In conjunction with the appropriate keys, this system will enable identification of most foodborne fungi to species level in just 7 days. For a few kinds of fungi, notably yeasts and xerophiles, subsequent growth under other more specialised conditions will be necessary. Finally, this book is dedicated to the general food microbiologist. May it help to restore equilibrium and assist in continued employment, when the quality assurance manager demands: ‘‘What is it?’’ . . . ‘‘How did it get in?’’ . . . ‘‘What does it do?’’ . . . ‘‘How do we get rid of it?’’ . . . and, worst of all . . . ‘‘Is it toxic?’’

Chapter 2

The Ecology of Fungal Food Spoilage

Food is not commonly regarded as an ecosystem, perhaps on the basis that it is not a ‘‘natural’’ system. Nevertheless an ecosystem it is and an important one, because food plants and the fungi that colonise their fruiting parts (seeds and fruit) have been co-evolving for millennia. The seed and nut caches of rodents have provided a niche for the development of storage fungi. Fallen fruit, as they go through the cycle of decay and desiccation, have provided substrate for a range of fungi. Humans have aided and abetted the development of food spoilage fungi through their vast and varied food stores. It can be argued, indeed, some rapidly evolving organisms, such as haploid asexual fungi, are moving into niches created by man’s exploitation of certain plants as food. Food by its very nature is expected to be nutritious: therefore, food is a rich habitat for microorganisms, in contrast with the great natural systems, soil, water and plants. Given the right physico-chemical conditions, only the most fastidious microorganisms are incapable of growth in foods, so that factors other than nutrients usually select for particular types of microbial populations. Perhaps the most important of these factors relates to the biological state of the food. Living foods, particularly fresh fruits, vegetables, and also grains and nuts before harvest, possess powerful defence mechanisms against microbial invasion. The study of the spoilage of such fresh foods is more properly a branch of plant pathology than food microbiology. The overriding factor determining spoilage of a fresh, living food is the ability of specific microorganisms to overcome defence mechanisms. Generally speaking, then, spoilage of

fresh foods is limited to particular species. Such specific relationships between fresh food and fungus are discussed in Chapter 11 and under particular species. Other kinds of foods are moribund, dormant or nonliving, and the factors which govern spoilage are physical and chemical. There are eight principal factors: (l) water activity; (2) hydrogen ion concentration; (3) temperature – of both processing and storage; (4) gas tension, specifically of oxygen and carbon dioxide; (5) consistency; (6) nutrient status; (7) specific solute effects; and (8) preservatives. Each will be discussed in turn below.

2.1 Water Activity Water availability in foods is most readily measured as water activity. Water activity (aw), is a physicochemical concept, introduced to microbiologists by Scott (1957), who showed that aw effectively quantified the relationship between moisture in foods and the ability of microorganisms to grow on them. Water activity is defined as a ratio: aw ¼ p=po ; where p is the partial pressure of water vapour in the test material and po is the saturation vapour pressure of pure water under the same conditions.

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_2, Ó Springer ScienceþBusiness Media, LLC 2009

3

4

2

Water activity is numerically equal to equilibrium relative humidity (ERH) expressed as a decimal. If a sample of food is held at constant temperature in a sealed enclosure until the water in the sample equilibrates with the water vapour in the enclosed air space (Fig. 2.1a), then aw ðfoodÞ ¼ ERH ðairÞ=100: Conversely, if the ERH of the air is controlled in a suitable way, as by a saturated salt solution, at equilibrium the aw of the food will be numerically equal to the generated ERH (Fig. 2.1b). In this way, aw can be experimentally controlled, and the relation of aw to moisture (the sorption isotherm) can be studied. For further information on water activity, its measurement and significance in foods see Duckworth (1975); Pitt (1975); Troller and Christian (1978); Rockland and Beuchat (1987). In many practical situations, aw is the dominant environmental factor governing food stability or spoilage. A knowledge of fungal water relations will then enable prediction both of the shelf life of foods and of potential spoilage fungi. Although the water relations of many fungi will be considered individually in later chapters, it is pertinent here to provide an overview. Like all other organisms, fungi are profoundly affected by the availability of water. On the aw scale, life as we know it exists over the range 0.9999þ to 0.60 (Table 2.1). Growth of animals is virtually confined to 1.0–0.99 aw; the permanent wilt point of mesophytic plants is near 0.98 aw; and most microorganisms cannot

Fig. 2.1 The concept of water activity (aw) (a) the relationship between aw and equilibrium relative humidity (ERH); (b) one method of controlling aw by means of a saturated salt solution, which generates a specific ERH at a specific constant temperature

The Ecology of Fungal Food Spoilage

grow below 0.95 aw. A few halophilic algae and bacteria can grow in saturated sodium chloride (0.75 aw), but are confined to salty environments. Ascomycetous fungi and conidial fungi of ascomycetous origin comprise most of the organisms capable of growth below 0.9 aw. Fungi capable of growth at low aw, in the presence of extraordinarily high solute concentrations both inside and out, must be ranked as among the most highly evolved organisms on earth. Even among the fungi, this evolutionary path must have been of the utmost complexity: the ability to grow at low aw is confined to only a handful of genera (Pitt, 1975). The degree of tolerance to low aw is most simply expressed in terms of the minimum aw at which germination and growth can occur. Fungi able to grow at low aw are termed xerophiles: one widely used definition is that a xerophile is a fungus able to grow below 0.85 aw under at least one set of environmental conditions (Pitt, 1975). Xerophilic fungi will be discussed in detail in Chapter 9. Information about the water relations of many fungi remains fragmentary, but where it is known it has been included in later chapters.

2.2 Hydrogen Ion Concentration At high water activities, fungi compete with bacteria as food spoilers. Here pH plays the decisive role. Bacteria flourish near neutral pH and fungi cannot compete unless some other factor, such as low water

2.3 Temperature

5

Table 2.1 Water activity and microbial water relations in perspectivea aw

Perspective

Foods

1.00

Blood, plant wilt point, seawater

0.95

Most bacteria

Vegetables meat, milk fruit Bread

0.90 0.85 0.80

Staphylococcus aureus

Ham Dry salami

0.75

Salt lake Halophiles

0.70

Jams Salt fish Fruit cake Confectionery Dried fruit Dry grains

Moulds

Yeasts

Basidiomycetes Most soil fungi Mucorales Fusarium Rhizopus, Cladosporium Aspergillus flavus Xerophilic Penicillia Xerophilic Aspergilli Wallemia Eurotium Chrysosporium Eurotium halophilicum

Basidiomycetes Most ascomycetes Zygosaccharomyces rouxii (salt) Zygosaccharomyces bailii Debaryomyces hansenii

0.65 Xeromyces bisporus Zygosaccharomyces rouxii (sugar) 0.60 DNA disordered a Modified from data of J.I. Pitt as reported by Brown (1974). Water activities shown for microorganisms approximate minima for growth reported in the literature.

activity or a preservative, renders the environment hostile to the bacteria. As pH is reduced below about 5, growth of bacteria becomes progressively less likely. Lactic acid bacteria are exceptional, as they remain competitive with fungi in some foods down to about pH 3.5. Most fungi are little affected by pH over a broad range, commonly 3–8 (Wheeler et al., 1991). Some conidial fungi are capable of growth down to pH 2, and yeasts down to pH 1.5. However, as pH moves away from the optimum, usually about pH 5, the effect of other growth limiting factors may become apparent when superimposed on pH. Figure 2.2 is an impression of the combined influence of pH and aw on microbial growth: few accurate data points exist and the diagram is schematic. For heat-processed foods, pH 4.5 is of course critical: heat processing to destroy the spores of Clostridium botulinum also destroys all fungal spores. In acid packs, below pH 4.5, less severe processes may permit survival of heat-resistant fungal spores (Section 2.3).

2.3 Temperature The influence of temperature in food preservation and spoilage has two separate facets: temperatures during processing and those existing during storage.

As noted above, heat-resistant fungal spores may survive pasteurising processes given to acid foods. Apart from a few important species, little information exists on the heat resistance of fungi. Much of the information that does exist must be interpreted with care, as heating menstrua and conditions can vary markedly, and these may profoundly affect heat resistance. High levels of sugars are generally protective (Beuchat and Toledo, 1977). Low pH and preservatives increase the effect of heat (Beuchat, 1981a, b; Rajashekhara et al., 2000) and also hinder resuscitation of damaged cells (Beuchat and Jones, 1978). Ascospores of filamentous fungi are more heat resistant than conidia (Pitt and Christian, 1970; Table 2.2). Although not strictly comparable, data of Put et al. (1976) indicate that the heat resistance of yeast ascospores and vegetative cells is of the same order as that of fungal conidia. Among the ascomycetous fungi, Byssochlamys species are notorious for spoiling heat processed fruit products (Olliver and Rendle, 1934; Richardson, 1965). The heat resistance of B. fulva ascospores varies markedly with isolate and heating conditions (Beuchat and Rice, 1979): a D value between 1 and 12 min at 908C (Bayne and Michener, 1979) and a z value of 6–7C8 (King et al., 1969) are practical working figures. The heat resistance of

6

2

The Ecology of Fungal Food Spoilage

Fig. 2.2 A schematic diagram showing the combined influence of water activity and pH on microbial growth

B. nivea ascospores is marginally lower (Beuchat and Rice, 1979; Kotzekidou, 1997a). Ascospores of Neosartorya fischeri have a similar heat resistance to those of Byssochlamys fulva, but have been reported less frequently as a cause of food spoilage. Heat resistant fungi are discussed further in Chapter 4. Food products may be stored at ambient temperatures, in which case prevention of spoilage relies on other parameters, or under refrigeration, where temperature is expected to play a preservative

role. Food frozen to –108C or below appears to be microbiologically stable, despite some reports of fungal growth at lower temperatures. The lowest temperatures for fungal growth are in the range –7 to 08C, for species of Fusarium, Cladosporium, Penicillium and Thamnidium (Pitt and Hocking, 1997). Nonsterile food stored at ca. 58C in domestic refrigerators, where conditions of high humidity prevail, will eventually be spoiled by fungi of these genera. At high aw and neutral pH, psychrophilic bacteria may also be important (mostly Pseudomonas species).

Table 2.2 Comparative heat resistance of ascospores and conidiaa Survivors (%) Fungus

Spore type

Initial viable count/ml

Ascospores 5.0  102 Conidia 7.3  102 Eurotium chevalieri Ascospores 1.0  103 Conidia 8.9  102 Xeromyces bisporus Ascospores 1.0  103 Aspergillus candidus Conidia 3.8  102 Wallemia sebi Conidia 7.1  102 a Heated at temperatures shown for 10 min. Data from Pitt and Christian (1970). Eurotium amstelodami

508C

608C

708C

93 107 103 128 93 102 42

85 0.3 62 0.1 30 0 0

3 0 21 0 0.3 0 0

2.4 Gas Tension

Thermophilic fungi, i.e. those which grow only at high temperatures, are rarely of significance in food spoilage. If overheating of commodities occurs, however, in situations such as damp grain, thermophiles can be a very serious problem. Thermotolerant fungi, i.e. species able to grow at both moderate and high temperatures, are of much greater significance. Aspergillus flavus and A. niger, able to grow between ca. 8 and 458C, are among the most destructive moulds known.

2.4 Gas Tension Food spoilage moulds, like almost all other filamentous fungi, have an absolute requirement for oxygen. However, many species appear to be efficient oxygen scavengers, so that the total amount of oxygen available, rather than the oxygen tension, determines growth. The concentration of oxygen dissolved in the substrate has a much greater influence on fungal growth than atmospheric oxygen tension (Miller and Golding, 1949). For example, Penicillium expansum grows virtually normally in 2.1% oxygen over its entire temperature range (Golding, 1945), and many other common food spoilage fungi are inhibited only slightly when grown in nitrogen atmospheres containing approximately 1.0% oxygen (Hocking, 1990). Paecilomyces variotii produced normal colonies at 258C under 650 mm of vacuum (Pitt, unpublished). Most food spoilage moulds appear to be sensitive to high levels of carbon dioxide, although there are notable exceptions. When maintained in an atmosphere of 80% carbon dioxide and 4.2% oxygen, Penicillium roqueforti still grew at 30% of the rate in air (Golding, 1945), provided that the temperature was above 208C. In 40% CO2 and 1% O2, P. roqueforti grew at almost 90% of the rate in air (Taniwaki et al., 2001a). Xeromyces bisporus has been reported to grow in similar levels of carbon dioxide (Dallyn and Everton, 1969). Byssochlamys species appear to be particularly tolerant of conditions of reduced oxygen and/or elevated carbon dioxide. Growth of Byssochlamys nivea was little affected by replacement of nitrogen in air by carbon dioxide, and growth in carbon dioxide–air mixtures was proportional only to

7

oxygen concentration, at least up to 90% carbon dioxide (Yates et al., 1967). Both Byssochlamys nivea and B. fulva were capable of growth in atmospheres containing 20, 40 or 60% carbon dioxide with less than 0.5% oxygen, but inhibition increased with increasing carbon dioxide concentration (Taniwaki et al., 2001a). Byssochlamys fulva is capable of growth in 0.27% oxygen, but not in its total absence (King et al., 1969). It is also capable of fermentation in fruit products, but presumably only if some oxygen is present. At least some species of Mucor, Rhizopus and Fusarium are able to grow and ferment in bottled liquid products and sometimes cause fermentative spoilage. Growth under these conditions may be yeast-like. Species of Mucor, Rhizopus and Amylomyces used as starter cultures in Asian fermented foods can grow under anaerobic conditions, demonstrated by growth in an anaerobe jar with a hydrogen and carbon dioxide generator (Hesseltine et al., 1985). Other authors have reported growth under anaerobic conditions of such fungi as Mucor species, Absidia spinosa, Geotrichum candidum, Fusarium oxysporum and F. solani (Stotzky and Goos, 1965; Curtis, 1969; Taniwaki, 1995). The yeast-like fungus Moniliella acetoabutans can cause fermentative spoilage under totally anaerobic conditions (Stolk and Dakin, 1966). As a generalisation, however, it is still correct to state that most food spoilage problems due to filamentous fungi occur under aerobic conditions, or at least where oxygen tension is appreciable, due to leakage or diffusion through packaging. In contrast, Saccharomyces species, Zygosaccharomyces species and other fermentative yeasts are capable of growth in the complete absence of oxygen. Indeed, S. cerevisiae and Z. bailii can continue fermentation under several atmospheres pressure of carbon dioxide. This property of S. cerevisiae has been harnessed by mankind for his own purposes, in the manufacture of bread and many kinds of fermented beverages. Z. bailii, on the other hand, is notorious for its ability to continue fermenting at reduced water activities in the presence of high levels of preservatives. Fermentation of juices and fruit concentrates may continue until carbon dioxide pressure causes container distortion or explosion. The closely related species Zygosaccharomyces rouxii is a xerophile and causes

8

spoilage of low-moisture liquid or packaged products such as fruit concentrates, jams and dried fruit. The difference in oxygen requirements between moulds and fermentative yeasts is one of the main factors determining the kind of spoilage a particular commodity will undergo.

2.5 Consistency Consistency, like gas tension, exerts considerable influence over the kind of spoilage to which a food is susceptible. Generally speaking, yeasts cause more obvious spoilage in liquid products, because single celled microorganisms are able to disperse more readily in liquids. Moreover, a liquid substrate is more likely to give rise to anaerobic conditions and fermentation is more readily seen in liquids. In contrast, filamentous fungi are assisted by a firm substrate, and ready access to oxygen. The foregoing is not intended to suggest that yeasts cannot spoil solid products nor moulds liquids: merely that all other factors being equal, fermentative yeasts have a competitive advantage in liquids and cause more obvious spoilage under these conditions.

2.6 Nutrient Status As noted in the preamble to this chapter, the nutrient status of most foods is adequate for the growth of any spoilage microorganism. Generally speaking, however, it appears that fungal metabolism is best suited to substrates high in carbohydrates, whereas bacteria are more likely to spoil proteinaceous foods. Lactobacilli are an exception. Most common mould species appear to be able to assimilate any food-derived carbon source with the exception of hydrocarbons and highly condensed polymers such as cellulose and lignin. Most moulds are equally indifferent to nitrogen source, using nitrate, ammonium ions or organic nitrogen sources with equal ease. Some species achieve only limited growth if amino acids or proteins must provide both carbon and nitrogen. A few isolates classified in Penicillium subgen. Biverticillium are unable to utilise nitrate (Pitt, 1979b).

2

The Ecology of Fungal Food Spoilage

Some xerophilic fungi are known to be more demanding. Ormerod (1967) showed that growth of Wallemia sebi was strongly stimulated by proline. Xerophilic Chrysosporium species and Xeromyces bisporus also require complex nutrients, but the factors involved have not been defined (Pitt, 1975). Yeasts are often fastidious. Many are unable to assimilate nitrate or complex carbohydrates; a few, Zygosaccharomyces bailii being an example, cannot grow with sucrose as a sole source of carbon. Some require vitamins. These factors limit to some extent the kinds of foods susceptible to spoilage by yeasts. A further point on nutrients in foods is worth making here. Certain foods (or nonfoods) lack nutrients essential for the growth of spoilage fungi. Addition of nutrient, for whatever reason, can turn a safe product into a costly failure. Two cases from our own experience illustrate this point, both involving spoilage by the preservativeresistant yeast Zygosaccharomyces bailii. In the first, a highly acceptable (and nutritious) carbonated beverage containing 25% fruit juice was eventually forced from the Australian market because it was impractical to prepare it free of occasional Z. bailii cells. Effective levels of preservative could not be added legally and pasteurisation damaged its flavour. Substitution of the fruit juice with artificial flavour and colour removed the nitrogen source for the yeast. A spoilage free product resulted, at the cost of any nutritional value and a great reduction in consumer acceptance. The other case concerned a popular water-ice confection, designed for home freezing. This confection contained sucrose as a sweetener and a preservative effective against yeasts utilising sucrose. One production season the manufacturer decided, for consumer appeal, to add glucose to the formulation. The glucose provided a carbon source for Zygosaccharomyces bailii, and as a result several months production, valued at hundreds of thousands of dollars, was lost due to fermentative spoilage.

2.7 Specific Solute Effects As stated earlier, microbial growth under conditions of reduced water availability is most satisfactorily described in terms of aw. However,

2.9 Conclusions: Food Preservation

the particular solutes present in foods can exert additional effects on the growth of fungi. Scott (1957) reported that Eurotium (Aspergillus) amstelodami grew 50% faster at its optimal aw (0.96) when aw was controlled by glucose rather than magnesium chloride, sodium chloride or glycerol. Pitt and Hocking (1977) showed a similar effect for Eurotium chevalieri and reported that the extreme xerophiles Chrysosporium fastidium and Xeromyces bisporus grew poorly if at all in media containing sodium chloride as the major solute. In contrast Pitt and Hocking (1977) and Hocking and Pitt (1979) showed that germination and growth of several species of Aspergillus and Penicillium was little affected when medium aw was controlled with glucose–fructose, glycerol or sodium chloride. Zygosaccharomyces rouxii, the second most xerophilic organism known, has been reported to grow down to 0.62 aw in fructose (von Schelhorn, 1950). Its minimum aw for growth in sodium chloride is reportedly much higher, 0.85 aw (Onishi, 1963). Some fungi are halophilic, being well adapted to salty environments such as salted fish. Basipetospora halophila and Polypaecilum pisce grow more rapidly in media containing NaCl as controlling solute (Andrews and Pitt, 1987; Wheeler et al., 1988c). Such fungi have been called halophilic xerophiles to distinguish them from obligately halophilic bacteria.

2.8 Preservatives Obviously, preservatives for use in foods must be safe for human consumption. Under this constraint, food technologists in most countries are limited to the use of weak acid preservatives: benzoic, sorbic, nitrous, sulphurous, acetic and propionic acids – or, less commonly, their esters. In the concentrations permitted by most food laws, these acids are useful only at pH levels up to their pKa plus one pH unit, because to be effective they must be present as the undissociated acid. For studies of the mechanism of action of weak acid preservatives see Warth (1977, 1991); Brul and Coote (1999); Stratford and Anslow (1998) and Stratford and Lambert (1999).

9

The use of chemical preservatives in foods is limited by law in most countries to relatively low levels and to specific foods. A few fungal species possess mechanisms of resistance to weak acid preservatives, the most notable being Zygosaccharomyces bailii. This yeast is capable of growth and fermentation in fruit-based cordials of pH 2.9–3, of 458C Brix and containing 800 mg/L of benzoic acid (Pitt and Hocking, 1997). The yeast-like fungus Moniliella acetoabutans can grow in the presence of 4% acetic acid and survive in 10% (Pitt and Hocking, 1997). Of the filamentous fungi, Penicillium roqueforti appears to be especially resistant to weak acid preservatives and this property has been suggested as a useful aid to isolation and identification (Engel and Teuber, 1978).

2.9 Conclusions: Food Preservation It is evident from the above discussion that the growth of fungi in a particular food is governed largely by a series of physical and chemical parameters, and definition of these can assist greatly in assessing the food’s stability. The situation in practice is made more complex by the fact that such factors frequently do not act independently, but synergistically. If two or more of the factors outlined above act simultaneously, the food may be safer than expected. This has been described by Leistner and Rodel (1976) as the ‘‘hurdle concept’’. ¨ This concept has been evaluated carefully for some commodities such as fermented sausages and is now widely exploited in the production of shelf stable bakery goods and acid sauces. For most fungi, knowledge remains meagre about the influence of the eight parameters discussed here on germination and growth. However, sufficient information is now available that some rationale for spoilage of specific commodities by certain fungi can be attempted, especially where one or two parameters are of overriding importance. This topic is considered in later chapters devoted to particular commodities.

“This page left intentionally blank.”

Chapter 3

Naming and Classifying Fungi

As with other living organisms, the name applied to any fungus is a binomial, a capitalised genus name followed by a lower case species name, both written in italics or underlined. The classification of organisms in genera and species was a concept introduced by Linneaus in 1753 and it is the keystone of biological science. It is as fundamental to the biologist as Arabic decimal numeration is to the mathematician. Here the analogy ends: the concept of ‘‘base 10’’ is rigorous; the concept of a species, fundamental as it is, is subjective and dependent on the knowledge and concepts of the biologist who described it.

3.1 Taxonomy and Nomenclature: Biosystematics Once biologists began to describe species and to assemble them into genera, questions about their relationships began to arise: is species x described by Jones in 1883 the same as species y described by Smith in 1942? Does species z, clearly distinct from x and y in some characters, belong to the same genus? The study of these relationships is termed taxonomy. Modern taxonomy is based on sound scientific principles, but still involves subjective judgment. When the decision is made that species x and species y are the same, however, the taxonomist must follow clearly established procedures in deciding which name must be used (‘‘has priority’’). The application of these procedures is termed nomenclature and, for fungi, plants and algae, is governed by the International Code of Botanical Nomenclature (ICBN). The ICBN is a relatively complex document of about 70 Articles dealing with all aspects of

correctly naming plants, algae and fungi. It is amended every 6 years by special sessions at each International Botanical Congress and is republished thereafter. The 17th version of the ICBN (the Vienna code) is the most recently published (McNeill et al., 2006). The ICBN impinges only indirectly on the work of the practicing mycologist or microbiologist. It is nevertheless of vital importance to the orderly naming of all plant life; to ignore the ICBN is to invite chaos. Where confusion arises over the correct name for a botanical species – a constant source of irritation to the nontaxonomist – it stems usually from one of three causes: indecision by, or disagreement among, taxonomists on what constitutes a particular species; incorrect application of the provisions of the ICBN; or ignorance of earlier literature. To return to our example, when species x and species y are seen to be the same, x has priority because it was published earlier; y becomes a synonym of x. Important synonyms are often listed after a name to aid the user of a taxonomy, and this procedure has been followed here. Through ignorance, the same species name may be used more than once, for example, Penicillium thomii Maire 1915 and P. thomii K.M. Zalessky 1927. The name P. thomii has been given to two quite different fungi. Clearly P. thomii Maire has priority; the later name is not valid. To avoid ambiguity, correct practice in scientific publication is to cite the author of a species at first mention, and before any formal description. The ICBN provides rules to govern change of genus name also. In our example, if species z is transferred to the genus to which species x and y belong, it retains its species name but takes the new

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_3, Ó Springer ScienceþBusiness Media, LLC 2009

11

12

3

Naming and Classifying Fungi

genus name. The original author of the name z is placed in brackets after the species name, followed by the name of the author who transferred it to the correct genus. For example, Citromyces glaber Wehmer 1893 became Penicillium glabrum (Wehmer) Westling 1893 on transfer to Penicillium by Westling in 1911. Note the use of Latinised names: glaber (masculine) became glabrum (neuter) to agree with the gender of the genus to which it was transferred. Further points on the use of the ICBN arise from this example. P. glabrum retains its date of original publication, and therefore takes priority over P. frequentans Westling 1911 if the two species are combined. When Raper and Thom (1949) combined the two species, a taxonomically correct decision, they retained the name P. frequentans, which was nomenclaturally incorrect, causing confusion when Subramanian (1971) and Pitt (1979b) took up the correct name. It is worth pointing out that the confusion in this and similar situations arose from Raper and Thom’s action in ignoring the provisions of the ICBN, not from that of later taxonomists who correctly interpreted it.

connection between the Ascomycetes, the fungi that produce sexual spores in sacks, and the Deuteromycetes, where spores are always asexual, has been known for a long time. However, molecular taxonomy has provided the fundamental assurance needed to make this change. From the point of view of the food mycologist, this is a mixed blessing. The demands of the molecular systematists may yet make the taxonomy of foodborne fungi even more complicated. The taxonomic system used here is believed to be both practical and in line with the current ‘‘best practice’’ of the nomenclaturalists. The hierarchical subdivisions in Kingdom Fungi of interest in the present context are shown below, using as examples three genera and species important in food spoilage:

3.2 Hierarchical Naming

Note that names of genera, species and varieties are italicised or underlined, while higher taxonomic ranks are not. Three subkingdoms of the kingdom Fungi include genera of significance in food spoilage. As indicated in the examples above, these are Zygomycotina, Ascomycotina and (much less commonly) Basidiomycotina. Fungi from each of these subkingdoms have quite distinct properties, shared with other genera and species from the same subkingdom. Unlike other texts, this book will not rely on initial recognition of a correct subkingdom before identification of genus and species can be undertaken. Nevertheless, identification of the subkingdom can provide valuable information about a fungus, so the principal properties of these three subkingdoms are described below.

A given biological entity, or taxon in modern terminology, can be given a hierarchy of names: a cluster of related species is grouped in a genus, of related genera in families, of families in orders, orders in classes, and classes in subkingdoms. Similarly a species can be divided into smaller entities: subspecies, varieties and formae speciales (a term usually reserved for plant pathogens). In most modern classifications, the fungi are ranked, like plants and animals, as a separate kingdom. Traditionally, fungi have been divided into several subkingdoms, based on spore type and some environmental considerations. Modern molecular methods have revolutionised this. Fungi have been shown to be more closely related to animals than plants, where traditional taxonomy has always placed them. Some of the so-called ‘‘lower fungi’’ have been shown not to be fungi of all (though mycologists will no doubt continue to study them). The most important change from the point of view of the food mycologist is the demise of the subkingdom Deuteromycotina, and its absorption (almost entirely) into the Ascomycotina. The

Kingdom Subkingdom Class Order Family Genus Species Variety

Fungi Zygomycotina Zygomycetes Mucorales Mucoraceae Rhizopus stolonifer

Fungi Ascomycotina Plectomycetes Eurotiales Trichocomaceae Eurotium chevalieri intermedius

Fungi Basidiomycotina Wallemiomycetes Wallemiales Wallemiaceae Wallemia sebi

3.3 Zygomycotina Most fungi within the subkingdom Zygomycotina belong to the class Zygomycetes. Fungi in this class possess three distinctive properties:

3.4 Ascomycotina

1. Rapid growth. Most isolates grow very rapidly, often filling a Petri dish of malt extract agar with loose mycelium in 2–4 days. 2. Nonseptate mycelium. Actively growing mycelia are without septa (cross walls) and are essentially unobstructed. This allows rapid movement of cell contents, termed ‘‘protoplasmic streaming’’, which can be seen readily by transmitted light under the binocular microscope. In wet mounts the absence of septa is usually obvious (Fig. 3.1a). 3. Reproduction by sporangiospores. The reproductive structure characteristic of Zygomycetes is the sporangiospore, an asexually produced spore which in genera of interest here is usually produced inside a sac, the sporangium, on the end of a long specialised hypha. Sporangiospores are produced very rapidly. From the food spoilage point of view, the outstanding properties of Zygomycetes are very rapid growth, especially in fresh foods of high water activity; inability to grow at low water activities (no Zygomycetes are xerophiles); and lack of resistance to heat and chemical treatments. From the food safety point of view, Zygomycetes have rarely been reported to produce mycotoxins.

3.4 Ascomycotina The subkingdom Ascomycotina is distinguished from Zygomycotina by a number of fundamental characters, the most conspicuous being the production of septate mycelium (Fig. 3.1b). Consequent on

13

this, growth of fungi in this subkingdom is usually slower than that of Zygomycetes, although there are some exceptions. Fungi in the subkingdom Ascomycotina, loosely called ‘‘ascomycetes’’, characteristically produce their reproductive structures, ascospores, within a sac called the ascus (plural, asci, Fig. 3.2a, b). In most fungi, nuclei normally exist in the haploid state. At one point in the ascomycete life cycle, diploid nuclei are produced by nuclear fusion, which may or may not be preceded by fusion of two mycelia. These nuclei undergo meiosis within the ascus, followed by a single mitotic division and then differentiation into eight haploid ascospores. In most genera relevant to this work, asci can be recognised in stained wet mounts by their shape, which is spherical to ellipsoidal and smoothly rounded; size, which is generally 8–15 mm in diameter; and the presence when maturity approaches of eight ascospores tightly packed within their walls. At maturity asci often rupture to release the ascospores, which are thick walled, highly refractile, and often strikingly ornamented (Fig. 3.2c, d). Two other characteristics of asci are significant: generally they mature slowly, after incubation for 10 days or more at 258C, and they are usually borne within a larger, macroscopic body, the general term for which is ascocarp. Genera of interest here usually produce asci and ascospores within a spherical, smooth-walled body, the cleistothecium (Fig. 3.3a), or a body with hyphal walls, the gymnothecium (Fig. 3.3b). Ascospores are highly condensed, refractile spores, which are often resistant to heat, pressure

Fig. 3.1 (a) Nonseptate mycelium of Syncephalastrum racemosum; (b) septate mycelium of Fusarium equiseti

14

3

Naming and Classifying Fungi

Fig. 3.2 Asci and ascocarps: (a) asci of Talaromyces species; (b) asci of Byssochlamys fulva; (c) ascospores of Eupenicillium alutaceum; (d) ascospores of Neosartorya quadricincta. Bars ¼ 5 mm

and chemicals. Almost all xerophilic fungi are ascomycetes. Besides their sexual spores, ascospores, ascomycetes commonly produce asexual spores. Formed after mitotic nuclear division, these spores are borne singly or in chains, in most genera of interest here from more or less specialised hyphal structures. The general term for this type of spore is conidium (plural, conidia), but other more specialised terms

exist for specific kinds of conidia. Along the evolutionary process, some Ascomycetes with welldeveloped asexual stages lost the ability to produce ascospores, and rely entirely on conidia for dispersal. Conidia, and the specialised hyphae from which they are borne, are astonishingly diverse in appearance. The size, shape and ornamentation of conidia and the complexity of the structures producing them

Fig. 3.3 (a) Cleistothecia of Eupenicillium; (b) gymnothecia of Talaromyces. SEM. Bars = 50 mm

3.7 Dual Nomenclature

provide the basis for classification of Ascomycetes that no longer produce the ascosporic (sexual) stage. Lacking ascospores, conidial fungi are not usually heat resistant, but conidia may be quite resistant to chemicals. Some conidial fungi are xerophilic.

3.5 Basidiomycotina The Subkingdom Basidiomycotina includes mushrooms, puffballs and the plant pathogenic rusts and smuts. Until recently it was not considered of any interest to the food mycologist. However, molecular studies indicate that the small brown species Wallemia sebi, long a curiosity because of its lack of resemblance to any other fungus, is a basidiomycete. It has no obvious phylogenetic affinity with any other genus and has now been classified in its own order, Wallemiales (Zalar et al., 2005). Only one other species of foodborne fungi, Trichosporonoides nigrescens Hocking and Pitt (1981), has a known affinity with this subkingdom.

3.6 The Ascomycete – Conidial Fungus Connection It was established more than a century ago that many fungal species carry the genetic information to produce both ascospores and conidia. These two kinds of spores are produced by different mechanisms and have different functions, so they are not always formed simultaneously. Not surprisingly, mycologists sometimes have given different generic and species names to a single fungus producing both an ascosporic and a conidial state. The usage of these names under the ICBN depends on the circumstances under which they were originally given. Some of these circumstances are discussed briefly below. The ascomycete state, now usually referred to as the teleomorph, is regarded by nomenclaturalists as the more important reproductive state, and the name applied to the teleomorph should be used when the ascomycete state is present. If the conidial state is also in evidence, the fungus is now a holomorph and is still correctly known by the teleomorph name. If the conidial state, known as the

15

anamorph, has a separate name, this strictly speaking applies to the conidial state. It should be used only when the ascomycete state is absent, or to refer specifically to the conidial state if the ascomycete is present. However, the reader is warned that some anamorphic names are, and will continue to be, in common use for holomorphic fungi. Under the Articles of the ICBN, a generic name originally given to an anamorphic or conidial fungus cannot be used for a teleomorphic or ascomycetous fungus. For example, the name Penicillium, originally given to an anamorphic fungus with no known teleomorph, cannot be used for the teleomorphs later found to be produced by other Penicillium species. Such teleomorphs are classified in the genera Eupenicillium or Talaromyces, depending on whether ascospores are produced in cleistothecia or gymnothecia. Correct species names for the ascomycetous and conidial states of a single holomorphic fungus may or may not be the same, depending both on the circumstance in which the names were originally given, and on later synonymy. For example, Eupenicillium ochrosalmoneum Scott and Stolk and Penicillium ochrosalmoneum Udagawa refer to the teleomorph and anamorph of a single fungus. Udagawa (1959) described the anamorph; the teleomorph was later found, in the same isolate, by Scott and Stolk (1967). On the other hand, the anamorph of Eupenicillium cinnamopurpureum Scott and Stolk (1967) is Penicillium phoeniceum van Beyma (1933), with P. cinnamopurpureum Abe ex Udagawa (1959) as a synonym. Scott and Stolk (1967) found a teleomorph in Udagawa’s P. cinnamopurpureum; Pitt (1979b) later showed that this species was a synonym of the earlier P. phoeniceum. E. cinnamopurpureum, the first name applied to the teleomorph, is unaffected by this change in the anamorph name. In passing, note that ‘‘Abe ex Udagawa’’ indicates invalid (incomplete) publication of this species by Abe, with validation later by Udagawa. The species dates from the year of validation.

3.7 Dual Nomenclature An important point here is that some isolates of Penicillium phoeniceum regularly produce the teleomorphic state Eupenicillium cinnamopurpureum, while

16

others, taxonomically indistinguishable, fail to produce a teleomorph at all. Because of this, it is essential to have a separate name for teleomorph and anamorph. The system of two names for a single fungus, known as dual nomenclature, has a place in the classification of fungi despite its apparent complexity. In the descriptions in later chapters, fungi for which both teleomorphs and anamorphs are known have both names listed. As noted above, if both states are found in a particular isolate, the teleomorph name is the more appropriate: to use that given to the anamorph is not incorrect, but this name is more sensibly applied to the conidial state only. Dual nomenclature would be relatively simple if the relationship between anamorph and teleomorph was always one to one. This is not the case. As has already been mentioned, species classified in Penicillium may produce teleomorphs in two genera, Eupenicillium and Talaromyces. On the other hand Talaromyces produces anamorphs in two genera, Penicillium and Paecilomyces. Aspergillus is the anamorph of eight or ten teleomorphic genera. Most teleomorph–anamorph relationships encountered in food mycology belong to the genera mentioned here. These relationships will be described where necessary under these particular genera.

3.8 Practical Classification of Fungi Fungi are classified in a vast array of orders, families, genera and species. Among natural organisms, the numbers of taxa of fungi are rivalled only by those of the flowering plants and insects. Estimates of fungal species range as high as 1.5 million; only 5% of this number have so far been described (Hawksworth, 1991). Many fungi are highly specialised. Some will grow only in particular environments such as soil or water; many are obligate parasites and require a specific host, such as a particular plant species, and will not grow in artificial culture; many grow only in association with plant roots. From the point of view of the food microbiologist, these kinds of fungi are irrelevant. In one sense, most fungi which spoil foods are also highly specialised, their speciality being the ability to obtain nutrients from, and hence grow on, dead, dormant or moribund plant material more or less regardless of source. The

3

Naming and Classifying Fungi

principal factors influencing food spoilage by fungi are physico-chemical and have already been outlined in Chapter 2. The point being made here is that food spoilage fungi are classified in just a few orders and a relative handful of genera. For this reason there is much to be said for food mycologists avoiding the use of a traditional, hierarchical classification as outlined above and employing a less formal approach to the identification of the fungi of interest to them. In the present work, this pragmatic approach has been followed as far as possible:

 The use of specialised terms has been kept to a 







minimum, while being cognisant of the need for clarity of expression. Hierarchical classification has been avoided as far as possible, consistent with retaining a logical approach to the presentation of fungi which are related or of similar appearance. Identification procedures used have been designed to be simple and comprehensible, avoiding the use of specialised equipment or procedures unavailable in the routine laboratory. To this end, identification of nearly all species included in this work is based entirely on inoculation of a single series of Petri dishes, incubation under carefully standardised conditions and examination by traditional light microscopy. A standard plating regimen has been used for the initial examination of all isolates (except yeasts), so that identification procedures can be carried out without foreknowledge of genus or even subkingdom. Cultural characters, which can be broadly defined as the application of microbiological techniques to mycology, have been used throughout.

The use of cultural characters has long been implicit in the study of fungi in pure culture on artificial substrates, especially in such genera as Aspergillus and Penicillium, genera of paramount importance in food spoilage. In Penicillium, cultural characters have been used as taxonomic criteria since the turn of the 20th century, but have assumed greater importance through the work of Pitt (1973, 1979b), who used the measurement of colony diameters, following incubation under standardised conditions, as a taxonomic criterion. The use of

3.8 Practical Classification of Fungi

pure culture techniques and growth data in fungal taxonomy is now widespread. Food microbiologists, the primary audience for this book, are familiar with cultural techniques and the use of a wide range of media and varied incubation conditions, so the authors make no apology for the taxonomic approach used in the present work. This approach is a logical extension of the system used by the first author in Penicillium taxonomy and which has been found to have a much broader applicability. In the field of mycology, different genera have been studied by many different people of varied backgrounds and for different reasons. Consequently, keys and descriptions have been based on a wide variety of media, often traditional formulations incorporating all sorts of natural products. This heterogeneity makes comparisons difficult

17

and adds unnecessary complexity to the task of the nonspecialist confronted with a range of fungal genera. The approach used here has been to examine every isolate (excluding yeasts) by a single system: inoculation onto a standard set of Petri dishes and examination of them culturally and microscopically after 7 days incubation. Most of the genera and species included in this book can be identified immediately, at that point. Only in exceptional cases has it been found necessary to reinoculate isolates onto a further set of media in order to complete identification. The exceptional fungi are first the xerophiles, many of which grow poorly if at all on the standard media, and, second, genera such as Fusarium and Trichoderma, in which some species cannot readily be differentiated on the standard regimen. Details of the techniques used are given in Chapter 4.

“This page left intentionally blank.”

Chapter 4

Methods for Isolation, Enumeration and Identification

This chapter describes techniques and media suitable for the enumeration, isolation and identification of fungi from foods. Some techniques are similar to those used in food bacteriology; others have been developed to meet the particular needs of food mycology. Most of the media have been specifically formulated for foodborne fungi. The approach taken here is designed to provide a systematic basis for the study of food mycology. In 1984 a group of about 30 of the world’s foremost scientists in food mycology met in Boston, Massachusetts, USA, to hear and discuss a wide range of presentations that explored many aspects of methodology in food mycology. Agreement was reached on broad issues and areas requiring further work pinpointed. The proceedings were published as ‘‘Methods for the Mycological Examination of Food’’ (King et al., 1986). At a second workshop, held in Baarn, the Netherlands, in 1990, results of a number of collaborative studies on media and methods were presented and some standardised protocols developed. The proceedings, published as ‘‘Modern Methods in Food Mycology’’ (Samson et al., 1992), provided a comprehensive overview of current thinking in this field. The working group which organised those two workshops was then formalised as the International Commission on Food Mycology (ICFM), a commission under the auspices of the Mycology Division of the International Union of Microbiological Societies (IUMS). ICFM is dedicated to international standardisation of methods in food mycology. Subsequent ICFM workshops were held in Copenhagen, Denmark (1994), Uppsala, Sweden (1998), Samsø, Denmark (2003) and Key West,

Florida (2007). Papers from the third and fourth workshops were published in the International Journal of Food Microbiology and the proceedings of the fifth (Samsø) workshop were published as ‘‘Advances in Food Mycology’’ (Hocking et al., 2006a). The methodology described below is based on recommendations from ICFM and represents current thinking within the food mycology community. However, no formal endorsement from ICFM is implied.

4.1 Sampling It must be emphasised at the outset that results from mycological assays of foods are only as good as the samples used. However, sampling is beyond the scope of this text. Excellent treatises on sampling plans for food bacteriological purposes have been produced by the International Commission on Microbiological Specifications for Foods (ICMSF, 1986, 2002) and are generally applicable to food mycology.

4.2 Enumeration Techniques Quantification of the growth of filamentous fungi is more difficult than for bacteria or yeasts. Vegetative growth consists of hyphae, which are not readily detached from the substrate and which survive blending poorly. When sporulation occurs,

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_4,  Springer ScienceþBusiness Media, LLC 2009

19

20

very high numbers of spores may be produced, causing sharp rises in viable counts, often without any great increase in biomass. The estimation of fungal growth or biomass is not easy, because no primary standard exists (such as cell numbers used for yeasts and bacteria). Although techniques for quantifying biomass have improved in recent years, most food laboratories continue to rely on viable counting (dilution plating) for detecting and quantifying fungal growth in foods. As well as dilution plating, a second standard method, known as direct plating, has been developed for estimating fungal numbers and growth in foods. Both methods are described in detail below. Techniques for biomass estimation will be discussed later.

4.2.1 Direct Plating Contributions at the international workshops mentioned above emphasised the use of direct plating as the preferred method for detecting, enumerating and isolating fungi from particulate foods such as grains and nuts. In direct plating, food particles are placed directly on solidified agar media. In most situations, particles should be surface disinfected before plating, as this removes the inevitable surface contamination arising from dust and other sources and permits recovery of the fungi actually growing in the particles. This process provides an effective measure of inherent mycological quality and permits assessment of the potential presence of mycotoxins as well. Surface disinfection should be omitted only where surface contaminants become part of the downstream mycoflora, for example, in grain intended for flour manufacture. Even here, surface disinfection before direct plating provides the most realistic appraisal of actual grain quality. Results from direct plating analyses are expressed as percentage infection of particles. The technique provides no direct indication of the extent of fungal invasion in individual particles. However, it is reasonable to assume that a high percentage infection is correlated with extensive invasion in the particles and a higher risk of mycotoxin occurrence.

4 Methods for Isolation, Enumeration and Identification

The standard protocol recommended by the ICFM (Hocking et al., 2006a, p. 344) is given below, with amplification where necessary. Surface disinfection. Surface disinfection is carried out by immersing particles in a chlorine solution. Household chlorine bleach, nominally 4–5% active chlorine, is effective. Dilute the chlorine 1 to 10 with water before use, to provide an approximately 0.4% solution. Immerse particles for 2 min, stirring occasionally, then drain the chlorine. Chlorine solutions are rapidly denatured by organic matter, so it is important to use a surplus of chlorine solution (10 times the volume of the particles) and to use the solution only once. This process is conveniently carried out in 250–500 ml beakers. Place 50 or more particles in the beaker and add chlorine. To dislodge air bubbles, immediately stir with a pair of forceps, leaving them in the solution, and preferably cover the beaker with a watch glass. The watch glass simplifies decanting of the chlorine, and the forceps, disinfected by the chlorine, may be used to plate the particles. Studies in our laboratory have shown that the treatment outlined here may be inadequate under some conditions. In commodities such as peanuts or maize where high levels of Aspergillus flavus or Penicillium species may be present, surface disinfection may be difficult. Here 2 min immersion in 70% ethanol followed by 2 min in 0.4% chlorine is recommended. Rinsing. After the chlorine is poured off, particles may be rinsed once with sterile water. Use a 1 min treatment, with stirring, then pour the water off. Again a watch glass should cover the beaker during this period, and the sterile forceps should be used for stirring. It is not clear whether rinsing fulfils any essential function. Early direct plating regimens used agents such as mercuric chloride for disinfection, and rinsing was essential to remove such toxic materials before they penetrated the particles too deeply. However, chlorine is effectively denatured by the particles and is believed to penetrate very little. In our opinion, the rinsing step can be omitted without loss of efficacy of the treatment, with savings in time and materials, and reduced risk of recontamination from the air. Plating. After disinfection and the optional rinse, particles should be plated onto solidified agar, at the rate of 6–20 particles per plate, depending on

4.2 Enumeration Techniques

particle size. Use the disinfected forceps. Plating should be carried out immediately: keep the watch glass on the beaker if this is not possible. Incubation. Incubate plates upright, under normal circumstances for 5 days at 258C. See more detailed notes below. Examination. After incubation, examine plates visually, count the numbers of infected particles and express results as a percentage. Differential counting of various genera is often possible. Correct choice of media, a stereomicroscope and experience will all assist in this process.

4.2.2 Dilution Plating Dilution plating is the appropriate method for mycological analysis of liquid or powdered foods. It is also suitable for grains intended for flour manufacture and other situations where total fungal contamination is relevant. Sample preparation. The two most common methods of sample preparation for dilution plating are stomaching and blending: stomaching is recommended by ICFM (King et al., 1986). The Colworth Stomacher (Sharpe and Jackson, 1972), or equivalent equipment (e.g. BagMixer1, Interscience, Saint Nom La Brete`che, France), is a very effective device for dispersing and separating fungi from finely divided materials such as flour and spices, and soft foods such as cheeses and meats, and its use is strongly recommended. Treatment time in the stomacher should be 2 min. Harder or particulate foods such as grains, nuts or dried foods, like dried vegetables, should be soaked before stomaching. Soaking times from 30 to 60 min are generally sufficient, but for extremely hard particles such as dried legumes, soaking for up to 3 h may be required. Comminution in a Waring Blender or similar machine is a suitable alternative for these types of samples and may give a more satisfactory homogenate. Blending times should not exceed 60 sec, as longer treatments may fragment mycelium into lengths too short to be viable or overheat the homogenate. The sample size used should be as large as possible. If a stomacher or BagMixer 400 is used, a sample size of 10–40 g is suitable.

21

Diluents. The recommended diluent is aqueous 0.1% peptone (Hocking et al., 2006a, p. 344), suitable for both filamentous fungi and yeasts. Saline solutions, phosphate buffer or distilled water are no longer recommended by ICFM as they may be deleterious to yeasts (Mian et al., 1997). The addition of a wetting agent such as polysorbate 80 (Tween 80) may be desirable for some products, but the natural wetting ability of the peptone is usually adequate. Special diluents may be necessary in some circumstances. If yeasts are to be enumerated from dried products or juice concentrates, the diluent should also contain 20–30% sucrose, glucose or glycerol, as the cells may be injured or be susceptible to osmotic shock. Dilution. Serial dilutions of fungi are carried out by the same procedures as those used in bacteriology, and the recommended dilution rate is 1:10 (=1+9). Fungal spores sediment more quickly than bacteria, so it is important to draw aliquots for dilution or plating as soon as possible, preferably within 1 min (Beuchat, 1992). Plating. Spread (surface) plating is recommended. When pour plates are used, fungi develop more slowly from beneath the agar surface and may be obscured by faster growing colonies from surface spores. Hence spread plating allows more uniform colony development, improves the accuracy of enumeration of the colonies and makes subsequent isolation of pure cultures easier. The optimum inoculum for surface plating is 0.1 ml. Best results will be obtained if plates are dried slightly before use. After adding the inoculum, spread it evenly over the agar surface with a sterile bent glass rod (‘‘hockey stick’’). Sterilise the rod by flaming it with ethanol before use. A plate spinner is a useful accessory. It is usually possible to enumerate plates with up to 150 colonies, but if a high proportion of rapidly growing fungi are present, the maximum number which can be distinguished with any accuracy will be lower. Because of this restriction on maximum numbers, it may be necessary on occasion to accept counts from plates with as few as 10–15 colonies. Clearly, such limitations on numbers per plate and the overgrowth of slow colonies will result in higher counting errors than are usually achieved with bacteria or yeasts. Enumerating yeasts is less difficult. In the absence of filamentous fungi, from 30 to 300 colonies per

22

plate can be counted and errors will be comparable with those to be expected in bacterial enumeration. Incubation. The standard incubation conditions are 258C for 5 days. However, other conditions may be more suitable in some circumstances (see notes below). Reporting results. As in bacteriology, results from dilution plating are expressed as viable counts per gram of sample. Note that such results are not directly comparable with those obtained from direct plating and may not offer a direct indication of the extent of fungal growth.

4.2.3 Incubation Conditions As noted above, the standard incubation conditions specified by ICFM are 258C for 5 days (Hocking et al., 2006a). Undoubtedly, 258C is the most suitable temperature for routine work in temperate to subtropical environments. Few if any common fungi are sensitive to this temperature, even those which grow readily under refrigeration. Higher temperatures are unacceptable in the temperate zone: 308C is close to the upper limit for growth of some important Penicillium species. In tropical regions, incubation at 308C is recommended as a more realistic temperature for enumerating fungi from commodities stored at ambient temperatures. In cool temperate regions such as Europe, 228C may be a more suitable incubation temperature. When used for fungi, Petri dishes should be stored upright. The principal reason is that some common fungi can shed large numbers of spores during handling, which in an inverted dish will be transferred to the lid. Reinversion of the Petri dish for inspection or removal of the lid may liberate spores into the air or onto benches and cause serious contamination problems.

4.3 Sampling Surfaces Methods are outlined below for directly sampling the mycoflora of surfaces of commodities such as fruits, meats, cheeses, salamis and dried fish and also packaging materials, machinery and walls.

4 Methods for Isolation, Enumeration and Identification

The techniques are based on those described by Langvad (1980) for studying the fungal flora of leaves. If samples are particulate, or can be cut up, sterile forceps can be used to press pieces of a suitable size (up to about 10 mm2) onto a suitable medium in a Petri dish. The sample is then removed, leaving an impression, and any spores or mycelium transferred will form colonies within a few days. This technique is known as press plating. For packaging materials such as cardboard, an alternative method is to cut a piece which will fit in a standard Petri dish. The dish is prepared by adding a sterile filter paper moistened with 10% glycerol and then placing a bent glass rod on it as a separator. After adding the sample, a thin layer of an appropriate agar medium is poured over its surface. To reduce evaporation, the dish should be sealed with Parafilm or a similar material or placed in a polyethylene bag before incubation at 258C for a few days. If contamination levels are not too heavy, the number and types of moulds present can be effectively estimated by this method. Colonies may be subcultured for identification. For sampling walls or other surfaces, or for nondestructive sampling, impressions may be taken using adhesive tape. Carefully handled, tape coming from the roll will be virtually sterile. Press a short length of tape firmly onto the surface to be sampled, adhesive side down, then transfer it, still with the same side down, onto a suitable growth medium. After 1–2 days incubation at 258C, the tape may be removed to allow development and sporulation of colonies. Surface sampling techniques for assessment of sanitation in food production and processing areas are discussed by Evancho et al. (2001). Surfaces may be sampled by using sterile swabs or by agar contact methods. The swab method involves rubbing a moistened sterile cotton swab over the test surface and placing the swab in a dilution bottle to be subsequently diluted and plated on appropriate media. Agar contact plates, also known as RODAC (replicate organism direct agar contact) plates, are restricted to use on smooth or semi-smooth flat surfaces. An alternative to agar contact plates is the agar slice technique, where agar is filled into a syringe-like apparatus or into an artificial sausage casing. The solidified agar can be pushed out onto

4.5 Isolation Techniques

the surface to be sampled, then the portion making contact sliced off with a sterile scalpel or wire and placed in a Petri dish for incubation.

4.4 Air Sampling Air sampling in the food processing environment is discussed by Evancho et al. (2001). The simplest method of air sampling is by sedimentation or settle plates. A Petri dish containing an appropriate agar medium is exposed to the atmosphere for a fixed period of time (usually 15–60 min), then closed and incubated at 258C (Samson et al., 2004a). This method can be useful in food production areas, as it gives a direct indication of the number and types of fungi likely to come into contact with exposed product. However, the settle plate technique lacks precision, and volumetric air sampling is a much more reliable indicator of air quality. A number of air sampling devices are commercially available. Of these, the Anderson sampler (either the two-stage or six-stage model; Anderson Instruments Inc, Atlanta, GA, USA) is probably the best, giving accurate and consistent results (Buttner and Stetzenbach, 1993). However, the Anderson sampler is expensive and requires mains power or a large battery for operation. In factory situations, the dry cell battery-operated Biotest

23

Hycon RCS and RCS High Flow centrifugal air samplers (Biotest, Solihull, UK) and the MAS-100 air sampler (Merck, Darmstadt, Germany) are more convenient, as they are small and readily portable. The Biotest RCS Plus sampler was reported to give comparable results to larger and more sophisticated machines, but its sampling efficiency gradually decreased for particle sizes below 4 mm (Benbough et al., 1993).

4.5 Isolation Techniques The term ‘‘isolation’’ is used here in its strict sense: the preparation of a pure culture, free from any contamination and ready for identification.

4.5.1 Yeasts Streaking techniques commonly used for bacterial purification are equally suitable for the isolation of yeasts (Fig. 4.1). A method widely used by yeast specialists is to disperse a portion of a colony in 2– 3 ml of sterile water, then streak a single loop of this suspension over the whole surface of a plate, moving the loop slowly down from top to bottom while simultaneously moving it rapidly across the plate

Fig. 4.1 Petri dishes of yeasts growing on malt extract agar, showing a streaking method suitable for producing isolated colonies

24

from side to side. After suitable incubation, wellseparated single colonies should appear in the lower half of the plate (Fig. 4.1). If all of these single colonies appear to be of similar size and appearance (taking into account the effect of crowding), the culture may be judged to be pure. Microscopic checks of some single colonies are also desirable. Disperse a needle point of cells from a colony in a drop of water, add a cover slip and examine by bright field illumination at about 400. Cell outlines will be clearly visible. Note that, unlike those of bacteria, yeast cell sizes often vary considerably in a pure preparation. Purity is indicated not so much by uniformity of cell size within a preparation as by similarity in microscopic cell appearance from colony to colony. When a culture is considered to be pure, streak it onto an appropriate slant.

4.5.2 Moulds Streaking techniques are ineffective for filamentous fungi and are not recommended. Isolation depends on picking a small sample of hyphae or spores – judged to be pure by eye, by hand lens or preferably under the stereomicroscope – and placing this sample on a fresh plate as a point inoculum. Purity is subsequently judged by uniformity in appearance of the colony which forms after incubation. The appearance of a mixed culture depends on the growth rates of the fungi present. If rates are diverse, a mixed culture is often indicated by a clump of dense hyphae at the inoculum point, surrounded by looser wefts of spreading hyphae. With fungi of approximately equal growth rates, mixtures are often indicated by colonies with sectoring growth: sectors will show differences in mycelial, spore or reverse colours or in radial growth rates. The simplest starting place for isolating fungi is an enumeration plate with well-separated colonies. Use a needle of platinum or nichrome, preferably cut to a chisel point with a pair of pliers, or a steel sewing needle. Sterilise it by heating, then plunge the tip into cold agar and leave until cool – with nichrome or steel this will require several seconds. With the tip of the cold, wet needle pick off a few spores or a tuft of mycelium – just enough to be

4 Methods for Isolation, Enumeration and Identification

visible – and inoculate a single point on a plate or slant. The same procedure can be applied to mixed cultures arising from direct plating or surface sampling techniques. It is advisable to keep notes on the appearance of the colony area sampled, as this will give an indication of whether the culture which grows up is the same as that intended to be isolated. It is generally easy to isolate rapidly growing fungi from those which grow more slowly. The outermost hyphal tips are usually free of contamination. The reverse process is often much more difficult. The isolation of slowly growing fungi in the presence of rapidly growing ‘‘weeds’’ often requires skill, patience and ingenuity. It is desirable to watch the point inocula daily over several days at least, because a particular stage in the life cycle may give some advantage. The slow colony may germinate more rapidly, or have a sector accessible to a needle, or spore more freely. The use of higher or lower incubation temperatures, or media of low aw or low nutrient status, or the addition of dichloran or rose bengal (see below) may all be of value in this process. Freeing fungi from bacteria, long considered to be a very difficult procedure, has been greatly simplified in recent years with the advent of media containing antibiotics. With use of the media recommended in the next section, bacterial contamination of cultures should be a rare event. When a pure colony is obtained, it should be inoculated onto a slant of an appropriate medium and incubated until ready for identification. Again, moulds should always be inoculated at a single point, preferably near the centre of the slant. This permits the best colony development and sporulation in most fungi.

4.5.3 Short Term Storage For short to medium-term storage, fungi are usually stored on slants, most commonly in McCartney or Universal bottles which have the advantages of being free standing and having caps that can be sealed to retard drying during storage. During incubation and until colonies are fully mature, however, caps must be kept loose on slants in bottles. Moulds

4.6 Choosing a Suitable Medium

25

require free access to oxygen for typical growth and sporulation. Oxygen starvation during growth will at best lead to retarded sporulation or at worst death of the culture. Long-term preservation of fungi is dealt with later in this chapter.

4.6 Choosing a Suitable Medium Food laboratories often rely on a single all-purpose medium to produce a ‘‘yeast and mould’’ count in everything from raw material to finished product. But just as the food bacteriologist uses selective media for particular purposes, so too food mycologists are developing a range of media suited to specific applications. It is plainly unrealistic to expect a single medium to answer all questions about mould and yeast contamination in all foods. The fungi that spoil meats or fresh vegetables are not the same as those that grow on dried fish. Although often used for this purpose, very dilute media such as potato dextrose agar are of little or no value for enumerating fungi from dried foods.

The most important division in types of enumeration media lies between those suitable for high water activity foods, such as eggs, meat, vegetables and dairy products, and those suited to the enumeration of fungi in dried foods. The most suitable media for dried foods depend on the type of food, the major categories being foods low in soluble solids such as cereals, high sugar foods such as confectionery and dried fruit, and salt foods. A second consideration lies in whether the primary interest is in moulds, or yeasts, or both; and a third concerns the presence or absence of preservatives. Finally, media are available for specific mycotoxigenic fungi, notably Aspergillus flavus and related species and Penicillium verrucosum plus P. viridicatum. An overview of media considered most suitable for particular purposes is given in Table 4.1. The table is derived from Pitt and Hocking (1997), together with recent recommendations from ICFM (Samson et al., 1992; Hocking et al., 2006a).

Table 4.1 Recommended media for fungal detection, enumeration and isolationa Type of food

Selecting for

Medium

Remarks

Fresh foods: milk and milk products, fruit, cheese, sea foods

Moulds Yeasts General

DRBC TGY, MEA, OGY DRBC

Blend (where necessary) and dilution plate

Freshly harvested grains, nuts

General Dematiaceous Hyphomycetes Fusarium Yeasts

DRBC DRBC, CZID CZID TGY, MEA, OGY

Direct plate Direct plate Direct plate Dilution plate

Fruit juices, fresh

Yeasts

TGY, MEA, OGY

Dilution plate

Fruit juices, preserved

Preservative resistant yeasts

TGYA, malt acetic agar

Dilution plate

Fruit juices, to be pasteurised, or pasteurised products

Heat resistant moulds

PDA, MEA

Special protocol

Fruit juice concentrates

Xerophilic yeasts

MY50G

Special diluents

Dried foods in general

General

DG18

Direct plate

Stored cereals, nuts

General Dematiaceous Hyphomycetes Fusarium

DG18 DRBC, CZID CZID

Direct plate Direct plate Direct plate

26

4 Methods for Isolation, Enumeration and Identification

Table 4.1 Recommended media for fungal detection, enumeration and isolationa (continued) Type of food

Selecting for

Medium

Remarks

Grain for milling into flour

General

DG18

Stomach or blend and dilution plate

Dried fruit, confectionery, chocolate, etc.

Xerophilic moulds and yeasts

MY50G

Direct plate

Fastidious xerophiles – in presence of Eurotium spp.

MY50G MY70GF

Direct plate Direct plate

General

DG18

Halophilic xerophiles

MY5-12, MY10-12

Direct plate or press plate Direct plate or press plate

Fungi producing aflatoxins

AFPA

Direct or dilution plate

DRYS

Direct or dilution plate

Salt foods, e.g. salt fish

General

General Fungi producing ochratoxins a For medium acronyms, see Section 4.6.

4.6.1 General Purpose Enumeration Media To be effective, a general purpose enumeration medium must fulfil several requirements (Pitt, 1986). As these are sometimes overlooked, they are listed here:

 to inhibit bacterial growth completely, without  

  

affecting growth of foodborne fungi (filamentous or yeasts); to be nutritionally adequate and support the growth of fastidious fungi; to suppress the growth of rapidly spreading fungi, especially the Mucorales, but not to prevent their growth entirely, so they too can be enumerated; to slow radial growth of all fungi, to permit counting of a reasonable number of colonies per plate, without inhibiting spore germination; to promote growth of relevant fungi; and to suppress growth of soil fungi or others generally irrelevant in food spoilage.

Fulfilling the above requirements necessitates the use of potent inhibitory compounds, and there is sometimes a fine line between inhibition of undesirable microorganisms and suppression of growth of those being sought. Modern fungal enumeration media rely on the use of antibiotics at neutral pH

for the inhibition of bacteria. Such media allow better recovery of moribund and sensitive fungi than the acidified media commonly used in the past. For many years rose bengal has been added to media to slow colony spread, while more recently 2,6-dichloro-4-nitroaniline (dichloran) has been added to inhibit rapidly spreading moulds. Many common spoilage fungi, Aspergillus and Penicillium species in particular, develop better on media with adequate nutrients. Low nutrient media of very high aw, such as potato dextrose agar, have lost favour because they are selective against some species in these genera. The media described below are considered to be the most satisfactory general purpose enumeration media available at this time (Hocking et al., 2006a). Formulations are given in the Media Appendix. Dichloran rose bengal chloramphenicol (DRBC) agar. DRBC (King et al., 1979, Pitt and Hocking, 1997) is recommended for both moulds and yeasts. It is particularly suited to fresh and high aw foods (Hocking et al., 1992). This medium contains both rose bengal (25 mg/kg) and dichloran (2 mg/kg), which restrict colony spreading without affecting spore germination unduly. Compact colonies allow crowded plates to be counted more accurately. This combination of inhibitors also effectively restricts the rampant growth of most of the common mucoraceous fungi such as Rhizopus and

4.6 Choosing a Suitable Medium

27

Fig. 4.2 Petri dishes of (a) DRBC and (b) RBC showing effective control of Rhizopus growth by rose bengal and dichloran in DRBC

Mucor (Fig. 4.2), although it does not completely control some other troublesome genera such as Trichoderma. In routine use, it is recommended that DRBC plates be incubated away from light at 258C for 5 days. Dichloran 18% glycerol (DG18) agar. Hocking and Pitt (1980) developed DG18 to be selective for xerophilic fungi from low moisture foods such as stored grains, nuts, flour and spices. DG18 was designed for enumeration of a range of nonfastidious xerophilic fungi and yeasts. However, it has been shown since that it supports growth of the common Aspergillus, Penicillium and Fusarium species, as well as most yeasts, and many other common foodborne fungi. DG18 can now be described as a general purpose medium with emphasis on enumeration of fungi from dried foods. DG18 is also recognised to be a very useful medium for enumeration of airborne fungi (Wu et al., 2000; Samson et al., 2004a). It is an effective inhibitor of Mucoraceous fungi, and bacteria are totally suppressed. However, growth of Eurotium species (previously known as ‘‘the Aspergillus glaucus group’’) is still somewhat too rapid, and colonies may have diffuse margins. The addition of detergent to DG18 has been reported to be an improvement in this respect (Beuchat and de Daza, 1992). Although DG18 is a satisfactory isolation medium for Eurotium species, it is not suitable for their

identification. Eurotium species are usually identified on Czapek yeast extract agar with 20% sucrose (CY20S), described later in this chapter. Other general purpose media. Under circumstances where rapidly spreading moulds do not cause problems, two alternative general purpose enumeration media are satisfactory. These are rose bengal chloramphenicol agar (RBC; Jarvis, 1973), from which DRBC was developed, and oxytetracycline glucose yeast extract agar (OGY; Mossel et al., 1970). OGY has been found to be very suitable for yeasts in the absence of moulds (Andrews, 1992a). Most of the media discussed above are available in ready to use dehydrated form from media suppliers such as Oxoid, Difco, BBL, etc.

4.6.2 Selective Isolation Media Although considerable progress has been made in the past 20 years, the formulation of selective media for foodborne fungi still requires a great deal of research. The availability of effective media can greatly simplify the isolation and identification of significant food spoilage and mycotoxigenic fungi. Most attention has been paid so far to the requirements of xerophilic fungi because of their failure to develop on standard high aw media. For mycotoxigenic fungi, satisfactory media exist only for

28

Aspergillus flavus and closely related species, Penicillium verrucosum and P. viridicatum, and the genus Fusarium. These media are considered below. Media for Aspergillus flavus and related species. Bothast and Fennell (1974) developed Aspergillus differential medium after finding that Aspergillus flavus and A. parasiticus produced conspicuous, diagnostic orange yellow colours in the colony reverses in the presence of an appropriate nitrogen source and ferric salts. Few other fungi produced a similar colouration. It was later shown that these species produce aspergillic acid or noraspergillic acid, which react with ferric ammonium citrate to form the colour complex (Assante et al., 1981). However, development of the orange yellow reverse pigment is not indicative of aflatoxin production. This medium was reformulated by Pitt et al. (1983) to produce Aspergillus flavus and parasiticus agar (AFPA). As well as more effective concentrations of the active ingredients, AFPA contains dichloran and chloramphenicol to inhibit spreading fungi and bacteria, respectively. When incubated at 308C for 42–48 h, colonies of Aspergillus flavus, A. parasiticus and A. nomius are distinguished by bright orange yellow reverse colours. Only A. niger can be a source of error: it grows as rapidly as A. flavus and sometimes produces a yellow, but not orange, reverse. After 48 h A. niger colonies begin production of their diagnostic black or dark brown heads, which provide a ready distinction from A. flavus. Prolonged incubation of AFPA, beyond 4 days, is not recommended because A. ochraceus and closely related species may also produce a yellow reverse after this time. AFPA is recommended for the detection and enumeration of potentially aflatoxigenic fungi in nuts, maize, spices and other commodities. Its advantages include rapidity, as 48 h incubation is usually sufficient, specificity, and simplicity, as little skill is required in interpreting results. In consequence, it can be a simple, routine guide to possible aflatoxin contamination (Pitt, 1984). This medium is also very effective for enumerating A. flavus in soils, where levels down to 5 spores per gram can be detected. Where soils carry a heavy bacterial load, doubling the quantity of chloramphenicol or adding other antibiotics may be of value. Detection of aflatoxin production using Coconut Cream Agar. Dyer and McCammon (1994) developed

4 Methods for Isolation, Enumeration and Identification

Coconut Cream agar (CCA) to detect aflatoxin production in Aspergillus flavus and A. parasiticus. CCA can be made using any brand of commercial canned coconut cream (available from Asian food stores in many places). Dilute 50:50 with water, add agar (1.5%) and autoclave. Inoculate solidified plates with up to four colonies (picked with a wet needle) and incubate at 308C for 5–7 days. Examine, reverse upmost, under long wave length UV light. Colonies producing aflatoxins will fluoresce bluish white or white, especially in the centres. Ignore fluorescence from the Petri dish itself. Use an uninoculated coconut cream agar plate as a control (Dyer and McCammon, 1994). Plates inoculated with known nontoxigenic and toxigenic strains are also useful controls. A. parasiticus isolates almost always produce aflatoxins. Media for fungi producing ochratoxin A. Although ochratoxin A was first described from Aspergillus ochraceus, recent molecular studies indicate that A. westerdijkiae is the major ochratoxin A producing species in Aspergillus Section Circumdati (Frisvad et al., 2004). Aspergillus carbonarius is also an important source of ochratoxin A, particularly in grape products (Abarca et al., 2004; Leong et al., 2006a and references therein). However, there are no selective or indicative media for these fungi. When DG18 is used as the isolation medium, selection of colonies with light brown (ochre) or black sporulation is a good starting point for detecting A. westerdjikiae or A. carbonarius respectively. Coconut cream agar (Dyer and McCammon, 1994) can also be used to screen for ochratoxin production in A. carbonarius and species in Aspergillus Section Circumdati (Heenan et al., 1998). Plates are best incubated at 258C rather than 308C for potentially ochratoxigenic species of Aspergillus. Penicillium verrucosum is the most important ochratoxin a producer in the genus Penicillium (Pitt, 1987), although P. nordicum can also produce this mycotoxin (Frisvad and Samson, 2004). Frisvad (1983) developed Dichloran rose bengal yeast extract sucrose agar (DRYS) for selective enumeration of P. verrucosum. P. nordicum, P. viridicatum and P. aurantiogriseum are also selected by DRYS. P. verrucosum and P. nordicum produce ochratoxin A, P. verrucosum also produces citrinin, while the latter two species produce xanthomegnin and viomellein. According to Frisvad (1983), P. verrucosum colonies on DRYS have a violet brown reverse, and

4.6 Choosing a Suitable Medium

the latter two species produce yellow colonies with a yellow reverse. The incubation regimen recommended by Frisvad (1983) is 7–8 days at 208C. Subsequently, Frisvad et al. (1992) developed dichloran yeast extract sucrose 18% glycerol agar (DYSG) on which P. verrucosum produces a red brown reverse. Because of its lower aw, DYSG inhibits rapidly growing fungi such as Rhizopus and Mucor more effectively than DRYES. In a study of 76 samples of wheat, rye and barley, Lund and Frisvad (2003) found that DYSG was more effective than DRYES for screening grain samples for the presence of P. verrucosum and potential ochratoxin A contamination. Media for Fusarium species. Dichloran chloramphenicol peptone agar (DCPA; Andrews and Pitt, 1986) can be used to isolate Fusarium species from grains and other substrates. The medium was developed from Nash-Snyder medium (Nash and Snyder, 1962) a medium for enumeration of Fusarium species from soils. DCPA uses a low level of dichloran as a substitute for the high level of pentachloronitrobenzene (PCNB), and chloramphenicol rather than the antibiotic mixture used by Nash and Snyder (1962). When Fusarium species are dominant, DCPA is effective for their isolation from grains, animal feeds and soil. However, DCPA has been found to be less effective in mixed populations. DCPA is a very useful medium for the identification of Fusarium species because it often induces the abundant formation of macroconidia (Hocking and Andrews, 1987). A much more stringent medium, effective for isolation of most Fusarium species occurring in foods, is Czapek iprodione dichloran agar (CZID; Abildgren et al., 1987). As well as dichloran, this medium contains the fungicide iprodione. CZID is highly selective for Fusarium species, and is probably the best medium for Fusarium enumeration and isolation. CZID is suitable for isolation of Fusarium species by direct plating of surface disinfected grains and other commodities, or dilution plating of homogeneous samples such as flour. Some questions remain concerning whether CZID may be too selective and not support growth of all foodborne Fusarium species. However, the common species all grow well. Castella´ et al. (1997) developed a Fusarium selective medium (MGA 2.5) using 2.5 mg/L malachite green as the selective agent. They reported that MGA 2.5 was more selective than Nash-Snyder medium as it did not allow development of colonies of other fungal genera.

29

Bragulat et al. (2004) compared the efficacy of MGA 2.5 medium with DCPA, CZID and several other Fusarium media, using pure cultures of twelve Fusarium species commonly found in foods as well as naturally contaminated samples. They reported that there was no statistically significant difference in colony counts of the Fusarium spp. tested, but that colonies on MGA 2.5 were smaller than the other media. MGA 2.5 did not allow growth of other fungi such as Zygomycetes and yeasts from naturally contaminated samples, thus providing better selectivity than the other media. Media for dematiaceous Hyphomycetes. Although designed primarily for Fusarium isolation, CZID has been found in our laboratory to be very useful for isolating dematiaceous Hyphomycetes, provided that iprodione is added at half the usual concentration, as full strength iprodione tends to restrict colony diameters too severely. Alternaria, Bipolaris, Curvularia, Stemphylium and Ulocladium species will grow and sporulate well when incubated for 5 days at 258C with a 12 h photoperiod. Drechslera species will grow but will not sporulate on this medium. DRBC is also of value for isolating these fungi, but some species do not sporulate readily on it. DCPA, originally developed for isolation of Fusarium, may also be used as an isolation medium for the dematiaceous Hyphomycete genera mentioned above. Alternaria, Curvularia and related genera usually grow rapidly and sporulate well on DCPA. However, fungi should not be maintained or stored on DCPA for more than two weeks, as ammonia is produced by aging cultures. Media for xerophilic fungi. Xerophilic fungi are of great importance in food spoilage, and hence media and techniques for their enumeration and isolation have received much attention. Xerophiles range from those which grow readily on normal media and which are only marginally xerophilic, such as many Aspergillus and Penicillium species, to those, such as Xeromyces bisporus, which will not grow at all on normal (high aw) media. It is not surprising that no single medium is suitable for quantitative estimation of all xerophilic fungi found causing food spoilage. As noted earlier, DG18 was developed as a general medium for xerophiles, and remains the medium of choice for this purpose. DG18 should be used in any general examination of the mycoflora of dried foods. Media for extreme xerophiles. Fungi discussed here include Xeromyces bisporus, xerophilic

30

Chrysosporium species, Eremascus species, Eurotium halophilicum and the halophilic xerophiles Polypaecilum pisce and Basipetospora halophila. These extreme xerophiles grow slowly if at all on DG18 and are quickly overgrown by rapidly spreading xerophiles such as common Eurotium species. Such species require special media and techniques. Since the early 1950s, studies on extremely xerophilic fungi have been carried out in our laboratory (Scott, 1957). A very wide variety of media have been used for their cultivation. The most effective of these, suitable for all except the halophilic xerophiles, is malt extract yeast extract 50% glucose agar (MY50G; Pitt and Hocking, 1997). Isolation techniques for extreme xerophiles. As well as special media, extreme xerophiles require special techniques for isolation. If a low aw commodity such as ground spice, dried fruit, fruit cake, confectionery or dried fish shows signs of white mould growth, it is likely that the fungus responsible is an extreme xerophile. Fungi of this kind are usually sensitive to diluents of high aw and often cannot be isolated by dilution plating. For direct plating, a convenient technique is to place small pieces of sample, without surface disinfection, onto a rich, low aw medium, such as MY50G. Alternatively, examine the food under the stereomicroscope, which will often provide useful information in any case, and use an inoculating needle to pick off pieces of mycelium or spores from the surface of the spoiled food. Place these pieces (three to six per plate) directly onto MY50G. Colonies should develop after 1–3 weeks incubation at 258C. Quite frequently extreme xerophiles will be present in pure culture and can be readily isolated. Examination with the stereomicroscope will usually indicate whether this is the case. Selection of growth which appears to cover the range of types seen will assist isolation of the principal fungi present. Direct sampling by the press plating technique described earlier in this chapter can be useful for isolating xerophiles. Sometimes extreme xerophiles are accompanied by Eurotium species, also capable of growth at very low aw. The latter are identifiable under the stereomicroscope by their Aspergillus heads. Under these circumstances, isolation of extreme xerophiles is more difficult. A medium of sufficiently low aw to slow the growth of Eurotium species is necessary.

4 Methods for Isolation, Enumeration and Identification

The most satisfactory medium is malt extract yeast extract 70% glucose fructose agar (MY70GF). MY70GF is of similar composition to MY50G, except that it contains equal parts of glucose and fructose to prevent crystallisation of the medium at the concentration used (70% w/w). It is made in a similar manner to MY50G. Growth of even the extreme xerophiles on MY70GF is extremely slow, and plates should be incubated for at least 4 weeks at 258C. Once growth is apparent, pick off small portions of colonies and transfer them to MY50G, to allow more rapid growth and sporulation. Media for halophilic xerophiles. Some xerophilic fungi from salted foods such as salt fish grow more rapidly on media containing NaCl and hence are correctly termed halophilic xerophiles. Malt extract yeast extract 5% salt 12% glucose agar (MY5-12) and malt extract yeast extract 10% salt 12% glucose agar (MY10-12) are suitable for these fungi. Techniques for enumeration and isolation are similar to those described above for other extreme xerophiles.

4.6.3 Techniques for Yeasts The simplest enumeration and growth medium for most food spoilage yeasts is malt extract agar (MEA). Although originally introduced as a growth medium for moulds, its rich nutritional status makes it very suitable for yeasts, and its relatively low pH (usually near 5.0) reduces the possibility of bacterial contamination. More recently, tryptone glucose yeast extract agar (TGY) has been recommended for enumeration of yeasts (Hocking et al., 1992, 2006a). Due to a higher glucose concentration (10%) and higher pH (5.5–6.0), this medium is more effective in recovery of stressed yeast cells, and colony development is usually faster than on MEA. However, its higher pH means that an antibiotic should be incorporated for enumeration of yeasts from food samples which may contain bacteria. Recommended antibiotics are chloramphenicol or oxytetracycline at the concentrations used in DRBC and OGY. Both MEA and TGY are suitable for enumeration of yeasts in products such as fruit juices, fruit purees and yoghurt, where moulds are usually present only in low numbers.

4.6 Choosing a Suitable Medium

If large numbers of moulds are present, often the case with solid products such as cheese, a general purpose enumeration medium such as DRBC or DG18 should be used. For circumstances where yeast enumeration remains difficult on media such as DRBC, a valuable technique is that of de Jong and Put (1980), which involves a 3-day anaerobic incubation. They recommended using pour plates or streak plates of mycophil agar (BBL Microbiological Systems, Cockeysville, Maryland, USA), but malt extract agar should be equally suitable. Antibiotics may be added if necessary. Plates are incubated at 258C for 3 days in an anaerobic jar, then taken out and incubated aerobically for another 2 days at 258C. As many spoilage yeasts are able to grow anaerobically, colonies will commence development without the possibility of overgrowth by moulds. This technique is unlikely to be suited to the enumeration of yeasts on surfaces of fresh fruit or vegetables, as yeasts which colonise these surfaces are often strict aerobes. Enrichment for yeasts in liquid products. In liquid food products, low numbers of yeasts may be difficult to detect, but may have a serious potential to cause spoilage. Enrichment techniques are the only satisfactory way of monitoring products in these circumstances. For products or raw materials free from suspended solids and of low viscosity, standard membrane filtration techniques are a satisfactory method for detecting low numbers of yeasts. The filter can be placed directly onto a suitable medium such as MEA or TGY and staining can be carried out subsequently. Centrifugation may also be used, but has the disadvantage that only relatively small volumes of product can be screened. If products or raw materials are viscous, of low aw, or contain pulps and cannot be filtered efficiently, other enrichment techniques are needed. In many cases, the best enrichment medium is the product itself, diluted 1:1 with sterile water. A 1:1 dilution increases the aw of juice concentrates or honey to a level which will allow growth of potential spoilage yeasts without causing a lethal osmotic shock to the cells. If the product contains preservative, dilution will lower the concentration and permit cells to grow. To detect low numbers of spoilage yeasts in cordials, fruit juice concentrates and similar materials, simply decant half the product from the container

31

and replace it with sterile water. Leave the cap loose, incubate at room temperature or 308C and watch for evidence of fermentation. Shaking the container daily will help to detect gases resulting from fermentation. Classical enrichment techniques used in bacteriology can also be used for yeasts. TGY broth has been used very successfully in our laboratory for enrichment of low numbers of yeasts in liquid products. Add 10 ml of product to 90 ml TGY broth and incubate at 258C for 3–4 days, or 308C for 2–3 days. Look for signs of fermentation and streak out onto TGY agar. Detection of preservative resistant yeasts. A few species of yeasts are able to grow in products containing preservatives such as sorbic, benzoic and acetic acids or sulphur dioxide. The most important of these is Zygosaccharomyces bailii. The simplest and most effective way to screen for preservative resistant yeasts is to spread or streak product onto plates of malt acetic agar (MAA), which is MEA with 0.5% acetic acid added (Pitt and Richardson, 1973) or TGY with 0.5% acetic acid added (TGYA, Hocking, 1996). MAA and TGYA are made by adding glacial (16 N) acetic acid to melted and tempered basal medium to give a final concentration of 0.5%. Mix and pour immediately. These media cannot be held molten for long periods or remelted because of their low pH (approximately 3.2 and 3.8 for MAA and TGYA, respectively). The acetic acid does not need sterilisation before use. MAA and TGYA are suitable media for monitoring raw materials, process lines and products containing preservatives for resistant yeasts. They are also effective for testing previously isolated yeasts for preservative resistance. Erickson (1993) developed a selective medium for Zygosaccharomyces bailii. Zygosaccharomyces bailii medium (ZBM) is based on Sabouraud dextrose agar amended with fructose, NaCl, tryptone, yeast extract and trypan blue dye, then acetic acid (0.5%) and potassium sorbate (0.01%) are added to make the medium selective. It is designed as a plating medium for detection of Z. bailii in acidified ingredients in conjunction with hydrophobic grid membrane filtration (Erickson, 1993). When compared with MAA and TGYA in an interlaboratory study, ZBM was found to be highly selective for Zygosaccharomyces bailii, to the exclusion of other important preservative resistant yeasts such as Schizosaccharomyces pombe and Pichia

32

membranaefaciens. In addition, recovery of Z. bailii cells sublethally injured by lyophilisation was significantly lower on ZBM than on TGYA or MAA (Hocking, 1996). Its high selectivity and complex formulation make ZBM unsuitable for routine laboratory use as a medium for detection of preservative resistant yeasts. Enrichment of preservative resistant yeasts. A technique capable of detecting yeast numbers as low as 1 cfu/ml within 4 days has been developed in our laboratory (Hocking et al., 1996). This method is particularly suitable for the detection of Zygosaccharomyces bailii in raw materials or finished product, but can also be used for the detection of Schizosaccharomyces pombe, Pichia membranaefaciens and other species of preservative resistant yeasts. The method involves a 2to 3-day enrichment step followed by a plating step with a further 2 days of incubation. Triplicate 20 ml tryptone glucose yeast extract (TGY) broths containing 0.5% acetic acid (TGYA) are each inoculated with 1 g or 1 ml of product and the broths incubated at 308C. After incubation for 48 and 72 h, 0.1 ml from each broth is spread plated onto TGY agar containing 0.5% acetic acid and the plates incubated at 308C. The detection time of the method is shortened by incubating broths and plates at 308C rather than the traditional temperature of 258C, as the optimum growth temperature for Zygosaccharomyces bailii is 30–328C (Jermini and Schmidt-Lorenz, 1987b). The sensitivity of the method is greatly increased by using triplicate broths instead of single or duplicate broths and by spread plating 0.1 ml from each broth instead of streaking a loopful onto TGYA agar. This method has been used to detect low numbers of cells of Zygosaccharomyces bailii in experimentally inoculated cordial syrup, mayonnaise, salad dressing and barbecue sauce and other preservative-resistant yeasts such as Schizosaccharomyces pombe, Pichia membranaefaciens and some preservative resistant strains of Saccharomyces cerevisiae. Yeasts of intermediate preservative resistance (e.g. Debaryomyces hansenii, Candida krusei and Torulaspora delbrueckii) can also be detected by this method. Better recoveries were obtained using TGY than a malt extract agar and broth system, possibly due to the fact that the pH of TGY broth + 0.5% acetic acid is 3.8, compared with pH 3.2 for MEA + 0.5% acetic acid, and there is a lower concentration of glucose (2%) in the ME

4 Methods for Isolation, Enumeration and Identification

system compared with 10% in TGY. Yeasts which are unable to grow in the presence of acetic acid or other weak acid preservatives (sorbic or benzoic acids and their salts) are not detected by this method.

4.6.4 Techniques for Heat-Resistant Fungi Heat resistant spoilage fungi, such as Byssochlamys, Talaromyces, Neosartorya and Eupenicillium species can be selectively isolated from fruit juices, pulps and concentrates by laboratory pasteurisation using various methods (Beuchat and Rice, 1979; Hocking and Pitt, 1984; Beuchat and Pitt, 2001; Houbraken and Samson, 2006). Three methods are described here: a plating method based on that of Murdock and Hatcher (1978), a direct incubation method and a filtration method for liquid samples such as liquid sugar. Plating method. If the sample to be tested is more concentrated than 358Brix, it should first be diluted 1:1 with 0.1% peptone or similar diluent. For very acid juices such as passionfruit, normally about pH 2.0, the pH should be adjusted to 3.5–4.0. Two 50 ml samples are taken for examination. Erlenmeyer flasks (250 ml) or polyethylene Stomacher bags may be used as heat penetration into these containers will be rapid. If using Stomacher bags, the tops should be heat sealed. If a heat sealer is not available, the tops may be folded over and secured with a clip and should not be fully immersed. The two samples are heated in a closed water bath at 808C for 30 min, then rapidly cooled. Each 50 ml sample is then mixed with an equal volume of double strength MEA distributed over four 150 mm Petri dishes. The Petri dishes are loosely sealed in a plastic bag to prevent drying and incubated at 308C for up to 30 days. Plates are examined weekly for growth. Most moulds will produce visible colonies within 10 days, but incubation for up to 30 days will allow for the possible presence of badly heat damaged spores, which may germinate very slowly. This long incubation time also allows most moulds to mature and sporulate, aiding their identification. The main problem associated with this technique is the possibility of aerial contamination of the plates with common mould spores, which will give false positive results. The growth of green Penicillium

4.6 Choosing a Suitable Medium

colonies, or colonies of common Aspergillus species such as A. flavus and A. niger, is a clear indication of contamination as these fungi are not heat resistant. To minimise this problem, plates should be poured in clean, still air or a Class 2 biohazard cabinet if possible. If a product contains large numbers of heat resistant bacterial spores (e.g. Bacillus species), antibiotics can be added to the agar. The addition of chloramphenicol (100 mg/l of medium) will prevent the growth of these bacteria. Direct incubation method. A more direct method used for screening fruit pulps and other semisolid materials avoids the problems of aerial contamination. Place approximately 30 ml of pulp in each of three or more flat bottles such as 100 ml medicine flats. Heat the bottles in the upright position for 30 min at 808C and cool, as described previously. The bottles of pulp can then be incubated directly, without opening and without the addition of agar. They should be incubated flat, allowing as large a surface area as possible, for up to 30 days at 308C. Any mould colonies which develop will need to be subcultured onto a suitable medium for identification. If containers such as Roux bottles are available, larger samples can be examined by this technique, but heating times must be increased. Bottle contents should reach at least 758C for 20 min when checked by a thermometer suspended near the centre of the pulp. For further details of the above methods see Hocking and Pitt (1984) or Beuchat and Pitt (2001). Filtration method for liquid sugars. This method permits the detection of very low numbers of cells in clear liquids such as liquid sugar. Sample size should be at least 100 g, taken after vigorously shaking the container from which the sample is drawn. Add 100 ml diluent (0.1% aqueous peptone) to 2  50 g samples and mix well to dissolve. Filter both samples sequentially through the same sterile 0.45 mm membrane filter. After both samples have passed through the filter, rinse the interior of the funnel with 3  20– 30 ml volumes of sterile diluent. Remove the filter from the filter holder using sterile forceps and place it in a sterile bottle or Stomacher bag. Add 10 ml diluent to the bottle or bag containing the filter and place in a water bath at 758C for 30 min. Ensure the sample is submerged in the water bath (weigh down if necessary). Cool rapidly to room temperature, shake well, then divide the 10 ml of diluent between three Petri dishes. Add a generous portion of MEA with

33

antibiotics to each plate, mix the agar and sample well, then let the plates solidify. Incubate at 308C for up to 30 days, examining weekly. Count colonies and report count per 100 g. This method was developed by BCN Research Laboratories, Rockford, Tennessee, USA.

4.6.5 Other Plating Techniques Three other techniques which were developed for counting bacteria have been applied to fungal enumeration. Spiral plate count. Zipkes et al. (1981) evaluated the application of the spiral plate procedure to the enumeration of yeasts and moulds. They compared this procedure with the traditional pour plate and streak plate methods and found that spiral plating gave a higher overall recovery and lower replicate plating errors than the other two methods. The medium they used was potato dextrose agar, but the technique should be no less efficient using the media recommended here. Automation of spiral plate counting was studied by Manninen et al. (1991). They compared counts of pure bacterial, yeast and mould cultures using standard plating methods and spiral plate counts determined both manually and with a laser colony detector. They concluded that counts were not significantly different except where large colonies (10–15 mm diameter) of Rhizopus oligosporus were enumerated. Other Rhizopus and Mucor species are also likely to interfere with this method unless a suitable plating medium (such as DRBC) is used. Alonso-Calleja et al. (2002) found that the spiral plate technique compared well with standard plate counting on OGYE agar for enumeration of yeasts and moulds in goat’s milk cheeses; however, Garcı´ a-Armesto et al. (2002) found the method unsuitable for enumeration of yeasts in raw ewe’s milk. Hydrophobic grid membrane filters. Membrane filters overprinted with a square hydrophobic grid have been developed for rapid enumeration of bacteria. The hydrophobic grid membrane filter TM (HGMF) system is marketed as ISO-Grid by Neogen (Lansing, MI, USA). The HGMF ‘‘count’’ is determined by a most probable number (MPN) calculation. Brodsky et al. (1982) applied the HGMF technique to counting yeasts and moulds in foods. They compared it with spread plating on

34

potato dextrose agar amended with antibiotics and found that the HGMF technique produced higher counts in 2 days than the traditional method did after 5 days. The HGMF method could find use for evaluating the quality of raw materials before incorporation into a product. For example, Erickson (1993) has proposed a method using HGMF in conjunction with a selective medium for detection of the preservative resistant yeast Zygosaccharomyces bailii in acidified ingredients. The HGMF method was collaboratively trialled by 20 laboratories using 6 naturally contaminated foods. Although there were some differences between counts obtained by HGMF and the reference method (5 day pour plate), the differences were not significant, and the HGMF method was adopted by the AOAC International (Entis, 1996). Spangenberg and Ingham (2000) found that the HGMF method gave equivalent results to DRBC for enumeration of yeasts and moulds in grated low moisture Mozzarella cheese. Use of HGMF requires special holders for the square membrane filters, and an automated counting system is necessary to take full advantage of the method. The number of colonies on an HGMF can be counted visually, but it is a relatively slow process. TM Petrifilm Yeast and Mould. Petrifilm YM (3 M Company, St. Paul, MN, USA) is a proprietary system for enumerating fungi on a layer of medium enclosed in a plastic film, which eliminates the use of Petri dishes. A collaborative study carried out by the AOAC (Knight et al., 1997) concluded that Petrifilm YM gave comparable results with the BAM method (USFDA, 1992) for yeast and mould counts in five different food types inoculated with a cocktail of several species of Aspergillus and Penicillium and two species of yeast. Consequently, Petrifilm YM was adopted first action by AOAC International. Other workers (Beuchat et al., 1990, 2007; Taniwaki et al., 2001b; Ferrati et al., 2005) have shown that Petrifilm YM performed satisfactorily compared with conventional cultural methods when tested on a range of foods. However, Petrifilm was not particularly effective in inhibiting the growth of spreading moulds such as Rhizopus and Mucor. In addition, subculturing colonies for identification was more difficult from Petrifilm YM than from traditional Petri dishes. Petrifilm YM has a high aw and we do not recommend it for analysis of foods of less than 0.95 aw.

4 Methods for Isolation, Enumeration and Identification

4.7 Estimation of Fungal Biomass A deficiency in all of the enumeration techniques which rely on culturing fungi is that the result is at best poorly correlated with growth or biomass. Biomass is usually regarded as the fundamental measure of fungal growth in biotechnology, but it is not easy to quantify under the conditions existing in foods. Mycelial dry weight is most commonly used as a biomass estimate, but its relationship to mycelial wet weight and to metabolism varies widely in foods, due to the great influence of aw on both of the latter parameters. Fungi growing at reduced aw can be expected to be more dense than at high aw due to increased concentrations of internal solutes, though this is exceptionally difficult to measure experimentally. The question of a satisfactory fundamental measure of fungal biomass remains unanswered. Despite these basic problems, several chemical and biochemical techniques are available to estimate the extent of fungal growth in commodities. These techniques rely either on some unique component of the fungus that is not found in other microorganisms or foods or on immunological or molecular techniques. Some are still in the developmental phase: the most important ones are described briefly here.

4.7.1 Chitin Chitin is a polymer of N-acetyl-D-glucosamine and is a major constituent of the walls of fungal spores and mycelium. It also occurs in the exoskeleton of insects but is not present in bacteria or in foods. Hence the chitin content of a food or raw material can provide an estimate of fungal contamination. Chitin is most effectively assayed by the method of Ride and Drysdale (1972). Alkaline hydrolysis of the food sample at 1308C causes partial depolymerisation of chitin to produce chitosan. Treatment with nitrous acid then causes partial solubilisation and deamination of glucosamine residues to produce 2,5-anhydromannose, which is estimated colorimetrically using 3-methyl-2-benzothiazolone hydrazone hydrochloride as the principal reagent. Alkaline hydrolysis is more readily accomplished at

4.7 Estimation of Fungal Biomass

1218C in an autoclave (Jarvis, 1977). Improved assay sensitivity was achieved by derivatisation of glucosamine and other products with o-phthalaldehyde, separation by high performance liquid chromatography and detection of fluorescent compounds with a spectrofluorimeter (Lin and Cousin, 1985). Ekblad and Nasholm (1996) also described an HPLC method which measured fluorescence of a 9-fluorenylmethylchloroformate derivative of glucosamine. The chitin assay remains rather complex and slow, usually requiring about 5 h. A number of studies have indicated that the chitin assay is a valuable technique for estimating the extent of fungal invasion in foods such as maize and soybeans (Donald and Mirocha, 1977), wheat (Nandi, 1978) and barley (Whipps and Lewis, 1980) to estimate mycorrhizal fungi and fungal pathogens in plant material and soil (Ekblad and Nasholm, 1996; Ekblad et al., 1998; Penman et al., 2000; Singh, 2005) and measure wood rotting fungi (Nilsson and Bjurman, 1998). Particular attention has been paid to the possibility of developing the chitin assay as a replacement for the Howard mould count for tomato products (Jarvis, 1977; Bishop et al., 1982; Cousin et al., 1984). The chitin assay has some shortcomings and has been severely criticised by some authors (e.g. Sharma et al., 1977). The relationship between dry weight and chitin content varies at least twofold for different food spoilage fungi (Cousin et al., 1984; Lin and Cousin, 1985; Cousin, 1996). Some foods contain naturally occurring amino sugars such as glucosamine and galactosamine, which should be removed by acetone extraction prior to hydrolysis (Whipps and Lewis, 1980). Products from rot-free tomatoes give positive glucosamine assays even after acetone extraction (Cousin et al., 1984) and chitin content does not increase proportionally with fungal growth (Sharma et al., 1977). Insect contamination of grain samples has been reported to produce grossly misleading results (Sharma et al., 1977), but the presence of fruit flies in tomatobased products was less serious (Lin and Cousin, 1985). Materials such as stored grains frequently contain insect fragments and need to be checked before chitin assays are attempted. Because of these difficulties, the use of chitin as a chemical assay for fungi in foods has largely been superseded by the ergosterol assay.

35

4.7.2 Ergosterol Ergosterol is the major sterol produced by fungi, but at most is a minor component of plant sterols (Weete, 1974). Ergosterol occurs as a component of fungal cell membranes, so is inherently likely to be correlated with hyphal growth and biomass. It is therefore a good candidate as a chemical for measuring fungal growth in foods and raw materials. Methodology for estimating ergosterol in cereals was developed by Seitz et al. (1977, 1979). Samples are blended with methanol, saponified with strong alkali, extracted with petroleum ether and fractionated by high pressure liquid chromatography. Ergosterol is detected by ultraviolet absorption, optimally at 282 nm, a wavelength at which other sterols exhibit little or no absorbance. For refinements to this methodology, see Newell et al. (1988). Liquid chromatography or gas chromatography with mass spectrometric detection (LC-MS; GC-MS) have also been used to estimate ergosterol (Headley et al., 2002; Dong et al., 2006; Varga et al., 2006). GC-MS in combination with nondiscriminating flash pyrolysis (Py-GC-MS) provides the advantages of little sample preparation and small sample size (Parsi and Gorecki, 2006). The ergosterol assay provides a useful method for quantifying fungal growth, so numerous studies have assessed the relationship between fungal growth and ergosterol production, using both liquid and solid substrates. Using liquid cultures, Zill et al. (1988) showed a correlation between ergosterol production, mycelial wet weight and mycelial protein in Fusarium graminearum. Matcham et al. (1985) reported that ergosterol correlated better with mycelial dry weight than chitin or laccase, a polyphenol oxidase. Variation exists, however. Torres et al. (1992) reported that Aspergillus ochraceus grown in liquid culture showed a threefold increase in ergosterol concentration in relation to mycelial dry weight as the culture aged from 2 to 26 days. Other reports (Matcham et al., 1985; Newell et al., 1987; Seitz et al., 1979) have indicated smaller variations, only one to twofold. Marfleet et al. (1991) showed that fungal biomass and ergosterol levels were correlated for three representative fungal species over a range of aw on solid substrates but not in liquid media. Nout et al. (1987) showed that the ergosterol content of Rhizopus oligosporus varied widely, from 2 to 24 mg/mg

36

mycelial mass, and varied with substrate, aeration and growth phase. The ergosterol content was low during the rapid growth phase but tended to increase, at times sharply, as growth slowed. Taniwaki (1995) demonstrated that in fungi growing in atmospheres low in O2 and with elevated CO2 levels, the ergosterol content of hyphae was significantly reduced. In atmospheres containing 60% CO2, ergosterol content per unit of hyphal length was up to six times less than in air. Growth medium also affected ergosterol concentrations: on average, seven foodborne fungal species grown on PDA produced more than twice as much ergosterol per unit hyphal length as when grown on CYA (Taniwaki, 1995; Taniwaki et al., 2006). However, there was a reasonable correlation between ergosterol and mycelium dry weight for seven of the eight species tested. Eurotium chevalieri was the exception: this species produced little ergosterol and appeared to produce several other sterols (Taniwaki et al., 2006). Marı´ n et al. (2005) examined 16 species of food spoilage fungi and concluded that ergosterol content and colony diameters were better correlated to fungal biomass than fungal counts were. Marı´ n et al. (2008) showed that, for 14 common food spoilage fungi, correlation coefficients between ergosterol and colony diameters were sufficiently significant over a range of aw values (0.95–0.85), pHs (5–7) and potassium sorbate concentrations (0.5–1.5%) for both parameters to be useful in growth modelling. Quantifying ergosterol production in foods has proved more difficult. Seitz et al. (1977) showed a good correlation between damage in rice grains and their ergosterol content, between ergosterol in wheat and rainfall during the growing season and between ergosterol content and fungal invasion in several sorghum hybrids. Matcham et al. (1985) reported good correlations between linear extension of Agaricus bisporus grown on rice grains and chitin, ergosterol and laccase production. Ergosterol content correlated with colony counts of fungi on wheat grains at 0.95 aw but not at 0.85 aw (Tothill et al., 1992). Using a stereomicroscope for visual examination, they concluded that sound grain contained up to 6 mg/g ergosterol, microscopically mouldy grain 7.5–10 mg/g and visibly mouldy grain more than 10 mg/g ergosterol. From studies on ergosterol levels, colony counts and mould growth in a variety of grain samples, Schnu¨rer and Jonsson (1992) concluded that ergosterol correlated with colony counts

4 Methods for Isolation, Enumeration and Identification

better on DG18 (r = 0.77) than on MEA (r = 0.69). Ergosterol levels of food grade wheat ranged from 2.4 to 2.8 mg/g dry weight, samples from field trials (of unspecified quality) from 3.0 to 5.6 mg/g and feed grains from 8 to 15 mg/g dry weight. After an extensive survey of ergosterol levels in Danish crops, Hansen and Pedersen (1991) concluded that the normal levels of ergosterol in barley were 7.6 – 2.8, wheat for bread making 5.0 – 1.5, rye for bread making 6.8 – 2.2, peas 2.2 – 2.7 and rapeseed 2.4 – 1.3 mg/g dry weight. Ochratoxin A in barley correlated well with ergosterol content and reached significant levels when ergosterol increased to 25 mg/g dry weight. However, aflatoxin B1 became detectable in cottonseed meal when ergosterol reached only 4 mg/g. ‘‘Burned’’ rapeseed, a measure of quality, became significant when ergosterol reached 1.4 mg/g dry weight. Lamper et al. (2000) found that ergosterol content correlated well (r = 0.87) with deoxynivalenol levels in wheat inoculated with Fusarium graminearum or F. culmorum. Moraes et al. (2003) found that there was good correlation between mould counts and ergosterol content of Brazilian maize (r = 0.94) but a poor correlation (r = 0.4) between ergosterol and aflatoxin content. Pietri et al. (2004) found significant correlation between ergosterol content of Italian maize and the major mycotoxins, fumonisin B1 (1995 crop) or zearalenone and deoxynivalenol (1996 crop). Ergosterol content correlated strongly with fat acidity values and germination ability of stored canola (Pronyk et al., 2006). These authors also noted that Penicillium and Aspergillus species contributed more to ergosterol than Eurotium species. Ergosterol levels in sound canola were between 1.46 and 1.67 mg/g, whereas levels above 2 mg/g indicated significant levels of spoilage (Pronyk et al., 2006). Karaca and Nas (2006) examined ergosterol content of dried figs and found good correlation (r = 0.92) between aflatoxin and ergosterol in reject figs which were fluorescent, but no significant correlation with patulin content. Kadakal et al. (2005) found good correlation (r = 0.98) between ergosterol and patulin in apple juice and that both patulin (r = 0.99) and ergosterol (r = 0.99) were linearly related to the proportion of decayed apples used to make the juice. Ergosterol has been used to assess mould growth in cheese with variable results (Pecchini, 1997; Taniwaki et al., 2001a).

4.7 Estimation of Fungal Biomass

Ergosterol content has also been investigated as an indicator of the mycological status of tomato products. Battilani et al. (1996) found a significant correlation between ergosterol, Howard mould count (HMC) and fungal growth, but with a high level of uncertainty. Kadakal et al. (2004) found a linear relationship between degree of decay in tomato pulp and HMC (r = 0.97) and ergosterol (r = 0.96) and concluded that ergosterol has the potential to be used in quality assessment of tomatoes. Sio et al. (2000) described an improved method for extraction of ergosterol from tomato products. Other applications of ergosterol as a measurement of fungal biomass include estimation of mould spores in indoor air and aerosols (Miller and Young, 1997; Robine et al., 2005; Lau et al., 2006), estimation of wood decay by fungi (Eikenes et al., 2005) and estimation of fungi in soil and wetlands (e.g. Headley et al., 2002; Zhao et al., 2005). The ergosterol assay is reported to have a high sensitivity and, in contrast to the chitin assay, requires only 1 h for completion (Seitz et al., 1979). Despite its limitations, it appears to be a useful indicator of fungal invasion of foods and to hold promise as a routine technique for quality control purposes.

4.7.3 Impedimetry and Conductimetry Metabolites produced by growth of microorganisms in liquid media alter the medium’s impedance and conductance. The use of changes in these properties as a measure of bacterial growth was suggested by Hadley and Senyk (1975) and was first applied to yeasts by Evans (1982) and to moulds by Jarvis et al. (1983). Most of the subsequent work on fungi has been carried out with yeasts, but the methodology is often applicable to moulds also. During the 1980s there were a number of studies aimed at optimising media for inducing detectable and reproducible changes in either conductance or capacitance during fungal growth (see Pitt and Hocking, 1997). Watson-Craik et al. (1989, 1990) studied 27 mould species on a wide range of both commercially available and specially prepared media and concluded that conductance and capacitance were both medium and species specific. The use of media high in ammonium ions and glucose, with added yeast extract and peptone, decreased the

37

influence of product variability and induced higher conductance changes (Owens et al., 1992). An impedimetric method for detection of heat resistant fungi in fruit juices was described by Nielsen (1992). The detection limit in artificially contaminated juices was one Neosartorya ascospore per millilitre, detectable in 100 h. Huang et al. (2003) reported an impedance method for detection of bacteria and fungi in bottled water which shortened the detection time from 5 days to 27.1 h for fungi and from 48 to 11.3 h for bacteria. Although impedimetry and conductimetry promised to be effective rapid methods when used under well-defined conditions for a specific purpose with a particular kind of food, the methodology does not appear to have been broadly taken up for food mycology applications.

4.7.4 Adenosine Triphosphate (ATP) ATP has also been suggested as a measure of microbial biomass because bioluminescence techniques provide a very sensitive assay (Jarvis et al., 1983). Provided that background levels of ATP in plant or other cells are very low, or that microorganisms can be effectively separated from such other materials, the method has some potential as a microbial assay. A good correlation was shown between ATP production and viable counts of six species of psychrotrophic yeasts grown in pure culture (Patel and Williams, 1985). The effective detection of low levels of yeasts in carbonated beverages by ATP has also been reported (LaRocco et al., 1985). However, living plant cells contain high levels of ATP and fungi are often very difficult to separate from food materials. Moreover, extraction of molecules from fungal cells is notoriously difficult, so this potential may be difficult to realise in food mycology. The most widespread application of ATP bioluminescence in the food industry is for monitoring hygiene of surfaces in food production facilities (Easter, 2007).

4.7.5 Fungal Volatiles Methods for detection and characterisation of fungal volatiles are finding increasing applications in

38

food mycology, for the detection of fungal deterioration of grain and other food products, for the identification of particular fungi and for assessing potential mycotoxin contamination. Over the last two decades, there has been rapid growth in the development of gas sensor arrays. Gas sensors are chemical sensors that produce an electronic signal which is used as input into a pattern recognition system in order to recognise different volatiles and odours. This integrated system of gas sensor and pattern recognition is often called an ‘‘electronic nose or e-nose’’ (Gardner and Bartlett, 1994; Schaller et al., 1998). Arrays comprising equipment to collect headspace volatiles, analysis by GC-MS or other sensitive analytical methods, coupled with computer analysis are also commonly used. Deterioration of stored grain results from a combination of chemical and biological changes, with changes due to fungal growth often predominant. Deterioration is marked by off-odour development, loss of germinability, caking, rancidity and sometimes mycotoxin development (Abramson et al., 1980). Fungi produce volatile chemicals during growth and particular chemicals may be associated with grain deterioration (Kaminski et al., 1974, 1975). Sinha et al. (1988) monitored production of 3-methyl-1-butanol, 1-octen-3-ol and 3-octanone in stored grain and found that the presence of these volatiles usually correlated with seed infection by Alternaria alternata, Eurotium repens, Aspergillus versicolor and several Penicillium species. Octenol and octanone in particular seemed to be associated with deterioration in grain due to fungal growth. Adamek et al. (1992) identified methylfuran, 2-methylpropanol and 3-methylbutanol as the most important metabolites from Eurotium amstelodami, Aspergillus flavus, Penicillium cyclopium and Fusarium culmorum growing on wheat. The volatiles produced by several Penicillium species were also studied. Borjesson et al. (1989) collected ¨ volatiles produced by several food spoilage fungi grown in pure culture on wheat grain and found that some compounds, especially 3-methyl-1-butanol, were produced early in fungal growth and could be used as an early warning of potential deterioration, before fungal growth became visible. The production of volatile metabolites by Penicillium species in grain correlated well with carbon dioxide production and ergosterol formation (Borjesson et al., 1990, 1992). The use ¨

4 Methods for Isolation, Enumeration and Identification

of volatile compound production as an indicator of mould deterioration in grains has been extensively assessed and reviewed (Kaminski and Wasowicz, 1991; Schnu¨rer et al., 1999; Magan and Evans, 2000; Paolesse et al., 2006; Balasubramanian et al., 2007). Fungal volatiles can be used to detect potential mycotoxin contamination, to discriminate between fungal species (Sunesson et al., 1995; Keshri et al., 1998) and even between toxigenic and non toxigenic strains of particular fungi (Sahgal et al., 2007). Karlshøj et al. (2007a) used an electronic nose to differentiate between closely related Penicillium species (P. camemberti, P. nordicum, P. paneum, P. carneum, P. roqueforti and P. expansum) from cheese. Volatile profiles can be used to predict mould spoilage in bakery products (Vinaixa et al., 2004; Marı´ n et al., 2007a) and to detect and differentiate between toxigenic and non toxigenic P. verrucosum strains in bakery products (Needham and Magan, 2003). Volatile profiles have also been used to differentiate between toxigenic and non toxigenic Fusarium strains (Keshri and Magan, 2000; Demyttenaere et al., 2004), to identify mycotoxins (aflatoxins, ochratoxin A and deoxynivalenol) in durum wheat (Tognon et al., 2005) and to detect and quantify ochratoxin A and deoxynivalenol in barley (Olsson et al., 2002). This technology has also been applied to predict the presence of P. expansum and patulin in apple products (Karlshøj et al., 2007b) and to detect and discriminate diseases of potato tubers (Kushalappa et al., 2002) and stemend rot and anthracnose in mangoes (Moalemiyan et al., 2006). Electronic nose technology has also been used for early detection of moulds in libraries and archives (Pinzari et al., 2004).

4.7.6 Immunological Techniques Cell wall proteins of fungi produce antigens, which can be detected by immunological methods. Some antigens are derived from components common to a wide range of fungi, and hence are indicative of general fungal growth, while others are genus or even species specific. A variety of methods have been developed to take advantage of these fungal antigens, although none is in widespread commercial use.

4.7 Estimation of Fungal Biomass

Enzyme-linked immunosorbent assay (ELISA). The preparation of antigens from three common foodborne fungi (Penicillium aurantiogriseum, Mucor racemosus and Fusarium oxysporum) was described by Notermans and Heuvelman (1985). Preparation of immunoglobulin antibodies against these antigens was followed by development of an ELISA assay. Fungi were detected in both unheated and heat processed foods by this method. Antigens were relatively genus specific: the M. racemosus antigens reacted with other Mucor and Rhizopus species, and the Penicillium antigen reacted with the Aspergillus species tested. It was subsequently shown that the Penicillium antigen reacted with 43 of 45 Penicillium species tested and that antigen production correlated with mycelial weight and was unaffected by culture conditions, medium, temperature and aw (Notermans et al., 1986). The Penicillium antigen also reacted with Aspergillus flavus and the level of antigen correlated with aflatoxin production (Notermans et al., 1986). ELISA techniques have also been studied as a potential replacement for the Howard mould count. Antigens from tomato moulds (Alternaria alternata, Geotrichum candidum and Rhizopus stolonifer) were used to produce an ELISA test sensitive to 1 mg/g of mould in tomato. A correlation was observed between antigen formation and mould added to tomato puree, while background interference was very low (Lin et al., 1986). The method was tested against a broader range of foods, with encouraging results (Lin and Cousin, 1987). Robertson and Patel (1989) improved the sensitivity of the method for tomato paste by using a polyclonal antibody against Botrytis cinerea, Mucor piriformis and Fusarium solani in addition to the three species used by Lin et al. (1986). ELISA based methodology has been reported for the detection of Fusarium species in corn (Meirelles et al., 2006), cornmeal (Iyer and Cousin, 2003) and grain (Rohde and Rabenstein, 2005). Correlations with other measures of fungal growth (ergosterol, colony count, mycotoxin levels) were variable. Aspergillus species have also been examined as targets for immunological detection because of their importance in mycotoxin contamination. ELISA detection of A. ochraceus in wheat (Lu et al., 1995), coffee powder, chilli powders and poultry feed (Anand and Rati, 2006) has been investigated, with reasonable correction with other parameters

39

(amount of ochratoxin A, r ¼ 0.93; percentage infection r ¼ 0.89) reported by Lu et al. (1995) but poorer correlation with colony forming units (r ¼ 68) and glucosamine content (r ¼ 0.64). Anand and Rati (2006) did not attempt to correlate their ELISA with external measurements of fungal biomass, but used an internal comparison method using freeze dried mycelium. Immunological detection of aflatoxigenic fungi has also been the subject of several studies, using antibodies raised to A. flavus or A. parasiticus (Candlish et al., 1997; Tsai and Yu, 1997, 1999; Yong and Cousin, 2001). Tsai and Yu (1999) reported good correlation between viable count and ELISA measurements, depending on substrate, with correlation coefficients varying from 0.96 for wheat to 0.86 for peanuts in artificially inoculated samples. ELISA tests for Botrytis and Monascus in foods (Cousin et al., 1990) and for the detection in rice of Penicillium islandicum (Dewey et al., 1990) and Humicola lanuginosa (Dewey et al., 1992) have been described. ELISA tests have also been applied to detection of yeasts in dairy products (Garcia et al., 2004) and fruit juice (Yoshida et al., 1996), moulds in paprika powder (Kisko et al., 1998) and moulds in agricultural commodities (Park et al., 2003) with varying success. Linfield et al. (1995) reported that polyclonal antibodies raised against Botrytis allii were useful in early detection of neck rot of onions caused by this pathogen. Immunochemical detection of fungi in food and feeds has been reviewed by Li et al. (2000). Latex agglutination. A different approach to the immunological detection of fungi in foods has been the coating of latex beads with antibodies and detection of agglutination of the beads in the presence of antigens (Kamphuis et al., 1989; Notermans and Kamphuis, 1992; Stynen et al., 1992). It was found that 0.8 mm latex beads coated with antibodies from the extracellular polysaccharide produced by Penicillium digitatum specifically detected Aspergillus and Penicillium species (Kamphuis et al., 1989). Detection limits were as low as 5–10 ng/ml of the purified antigen. Commercially produced latex agglutination tests were of value for screening the mycological quality of grains and processed foods (Braendlin and Cox, 1992; van der Horst et al., 1992), although one kit performed poorly in detection of mould in tomato products (van der Horst et al., 1992).

40

Schwabe et al. (1992) compared the latex agglutination assay with ergosterol production for detection of Penicillium, Aspergillus and Fusarium species in pure culture. They concluded that the two methods were comparable for Penicillium and Aspergillus but that ergosterol was more sensitive for Fusarium. In food samples, both the latex agglutination test and ergosterol were effective means of detecting mould growth, but no clear correlation existed in values obtained by the two methods. Kesari et al. (2004) used a latex agglutination test to detect teliospores of Karnal bunt (Tilletia indica) in single grains of wheat and were able to detect a few as 750 teliospores, which they reported as suitable for single seed analysis. Fluorescent antibody techniques. Fluorescent antibody techniques have also been used directly for the detection of mould in foods. Warnock (1971) detected Penicillium aurantiogriseum in barley by this method, while Robertson et al. (1988) used antisera from five fungi to visualise moulds and simplify their detection in the Howard mould count technique.

4.7.7 Molecular Methods The first techniques for detecting DNA sequences using specific probes were devised more than 30 years ago (Southern, 1975). With the development of the polymerase chain reaction (PCR), gene cloning and oligonucleotide synthesis, DNA sequences can now be prepared in large quantities for use in probes. Depending on its role in the genome, DNA may be specific at almost any taxonomic level. Molecular methods can be used in both detection and identification of fungi, and real-time PCR methods even allow quantitative detection. This field of research continues to develop very rapidly, and significant advances may be expected. Producing a broadly based or even strain specific DNA probe is theoretically possible for any organism. Full genome sequences are available for a number of important foodborne fungi, including several Aspergillus species (A. nidulans, A. fumigatus, A. oryzae, A. flavus, A. niger, A. terreus, A. clavatus and Neosartorya fischeri), Botrytis cinerea, Chaetomium globosum, Fusarium graminearum, F. oxysporum, F. verticillioides, Rhizopus oryzae and several yeast species (Saccharomyces

4 Methods for Isolation, Enumeration and Identification

cerevisiae, Candida albicans, C. guilliermondii, C. lusitaniae, C. tropicalis, Schizosaccharomyces japonicus and Sch. pombe) (Broad Institute, 2008). Penicillium species are noticeably absent from the list, although the genome of the pathogenic species P. marneffei has recently been sequenced (JCVI, 2008). There have been rapid advances in molecular methods for the detection of mycotoxigenic fungi. Probes have been developed that target genes in the mycotoxin biosynthetic or regulatory pathways for many species of Aspergillus, Penicillium and Fusarium, enabling these fungi to be detected and/or quantified in grains, grapes, apples, coffee and other food matrices (Edwards et al., 2002; Paterson, 2006; Geisen, 2007; Niessen, 2007a, b; Rossi et al., 2007). Molecular methods have also been developed for the detection of spoilage fungi, particularly yeasts, e.g. Brettanomyces/Dekkera in grapes and wine (Phister and Mills, 2003; Agnolucci et al., 2007; Hayashi et al., 2007), Hanseniaspora in wine (Phister et al., 2007); Zygosaccharomyces bailii in wine and fruit juices (Rawsthorne and Phister, 2006) and Kluyveromyces marxianus in yoghurt (Mayoral et al., 2006). More general techniques have been described for detection of yeasts in juices (Casey and Dobson, 2004; Ros-Chumillas et al., 2007), milk, yoghurt and cheese (Cocolin et al., 2002; Bleve et al., 2003; Garcı´ a et al., 2004; Lopandic et al., 2006; Gente et al., 2007), sourdough fermentation (Meroth et al., 2003), table olives (Arroyo-Lopez et al., 2006) and vacuum packaged ham (Sanz et al., 2005). PCR based methods have also been described for detection of moulds in orange juice (Wan et al., 2006), Alternaria in cereal grains (Zur et al., 2002) and Botrytis in onion seeds (Walcott et al., 2004). As gene sequencing has become more readily available and cheaper, along with freely accessible databases such as GenBank (http://www.ncbi.nlm.nih.gov/Genbank/), sequence analysis and comparison is becoming a routine method for identification of yeasts and filamentous fungi. For yeasts, the most common target region is the D1/D2 domain of the 26S rDNA (Kurtzman et al., 2003), but the internal transcribed spacer (ITS) region of ribosomal DNA (rDNA) may also be used (van der Vossen et al., 2003; Pulvirenti and Giudici, 2003; Solieri et al., 2006; Villa-Carvajal et al., 2006).

4.8 Identification Media and Methods

Sequencing of the ITS region along with ‘‘housekeeping genes’’ such as calmodulin, b-tubulin and elongation factor 1-a is now commonly applied for purposes of identification and phylogenetic analysis of important food spoilage and mycotoxigenic fungi in the genera Aspergillus (Varga, 2006; Geiser et al., 2007; Peterson, 2008), Penicillium (Peterson, 2004, 2006; Samson et al., 2004b; Wang and Zhuang, 2007; Serra et al., 2008) and Fusarium (O’Donnell et al., 2004; Scott and Chakraborty, 2006; Leslie et al., 2007). Despite the apparent power of molecular techniques, they need to be applied with some caution, particularly when comparing DNA sequences with those in the publicly available databases to identify yeast or mould isolates. Correct identification relies on the database sequences having the correct name attached to them by the depositor, which is not always the case. If the identification makes sense, the percent homology is 98% or greater and the number of base pairs on which the homology is scored is high, then the answer is probably correct.

4.8 Identification Media and Methods 4.8.1 Standard Methodology The identification keys in this book are based primarily on the standardised procedure described for the identification of Penicillium species by Pitt (1979b). Cultures are grown for 7 days on three standard media at 258C, and on one of these at 5 and 378C also. The three Media are Czapek yeast extract agar (CYA; Pitt, 1973) used at all three temperatures; malt extract agar (MEA; Raper and Thom, 1949) and 25% glycerol nitrate agar (G25N; Pitt, 1973). Their formulae are given in the Media Appendix. Preparation time of CYA and G25N is reduced by the use of Czapek concentrate (Pitt, 1973), which is added to the media at the rate of 1% of the aqueous portion. As media ingredients have become more purified in recent years, difficulties with extent and colour of sporulation on CYA have been encountered, especially with some Penicillium species. To overcome this problem, Czapek concentrate has been reformulated (Pitt, 2000) by the inclusion of traces of zinc and copper (Smith, 1949) (see Media Appendix).

41

4.8.2 Plating Regimen As noted above, cultures for identification are grown on three media and at three temperatures. Maximum efficiency in time, incubator space and materials is achieved by inoculating two cultures on a single Petri dish of G25N, and at 5 and 378C, as shown in Fig. 4.3. Cultures are rarely mutually inhibitory under these conditions, although overgrowth of one culture by another is occasionally a problem at 378C. Standard sized Petri dishes (90–100 mm) are used, except that at 58C, smaller plates (50–60 mm in diameter) can be an advantage. The smaller size is easier to examine under the low power microscope. All plates are incubated for a standard time of 7 days. Plates incubated at 378C should be enclosed in polyethylene bags to prevent evaporation and drying of the medium. Unless the humidity is very low, plates at 258C will not dry excessively in 7 days. If no 58C incubator is available, use a polyethylene food container or insulated box in a household refrigerator. The box should be equipped with a thermometer, and its location moved by trial and error until a place with a 58C average temperature is located. Temperatures at 5 and 378C should ideally be –0.58C and be checked frequently; at 258C, –28C control is adequate (Okuda, 1994).

4.8.3 Inoculation As shown in Fig. 4.3, Petri dishes of CYA and MEA for incubation at 258C are inoculated with a single culture at three points, equidistant from the centre and the edge of the plate and from each other. Plates of the other media are inoculated with two points per culture, as illustrated. With some fungi, especially Penicillium and Aspergillus, it is important to minimise colonies from stray spores. The most satisfactory technique is to inoculate plates with spores suspended in semisolid agar (Pitt, 1979b). Dispense 0.2–0.4 ml of melted agar (0.2%) and detergent (0.05%), such as polysorbitan 80 (Tween 80), in small vials and sterilise. To use, add a needle point of spores and mycelium to a vial and mix slightly. Then, before flaming the needle, use it to stab inoculate the 58C plate;

42

4 Methods for Isolation, Enumeration and Identification

Fig. 4.3 Schematic of regimen used for culturing fungal isolates for identification

residual spores on the needle make a good inoculum. Next, take a sterile loop, mix the vial contents thoroughly and inoculate the standard plates. Used vials can be sterilised by steaming and reused several times before being washed or discarded.

4.8.4 Additional Media and Methods The plating regimen outlined above can be used to identify most of the fungi described in subsequent chapters of this book. Some exceptions exist, because certain genera either grow poorly or fail to sporulate on the standard media. As noted earlier in this chapter, fastidious xerophiles are identified on MY50G agar. Eurotium species, traditionally identified on Czapek agar with 20% sucrose, are identified here on Czapek yeast extract agar with 20% sucrose (CY20S; see Appendix). Trichoderma species are best identified on potato dextrose agar (PDA) after a relatively short incubation time (3–4 days), as the structures tend to autolyse as cultures mature. Penicillium subgenus Penicillium. Many species classified in subgenus Penicillium are morphologically similar, and identification using traditional morphological techniques remains difficult. These

Penicillia are very common in foods, and many produce mycotoxins, so correct identification is often critical. Species within subgenus Penicillium fall into two groups: those with an affinity for proteinaceous foods and those which grow more vigorously in foods high in carbohydrate. Frisvad (1981, 1985) introduced creatine sucrose agar (CREA) to permit differentiation between these two groups. Creatine (as the sole nitrogen source) permitted growth of the former group while inhibiting the latter. Incorporation of bromocresol purple enabled visualisation of pH changes, either acid or alkaline, depending on the creatine or sucrose metabolism of a particular species. However, discrimination between positive and negative responses was not always clear, and the main tabulation of species reactions to CREA (Table 1 in Frisvad, 1985) was difficult to interpret. Frisvad (1993) subsequently produced a number of variations of CREA, including acid and neutral pH formulations and substitution of sucrose with fructose or lactose. Although the merits of each formulation were discussed, no firm recommendation resulted from this exercise. Pitt (1993a) modified Frisvad’s strongly alkaline CREA medium by studying several sucrose and creatine concentrations over a wide pH range. The result was neutral creatine sucrose agar (CSN), a medium producing eight different reactions among

4.8 Identification Media and Methods

the 20 Penicillium subgenus Penicillium species tested. When included in the normal plating regimen for identification of Penicillium cultures from the subgenus Penicillium, CSN provides a very useful aid to distinguishing between these difficult and closely related species. See Chapter 7 for details of use of CSN, its reactions and interpretation. Its formulation is given in the Media Appendix. Dematiaceous Hyphomycetes. The natural habitats for many dematiaceous Hyphomycetes are plants or plant material, so finding suitable laboratory media and conditions to induce typical sporulation can be difficult. Alternaria, Curvularia, Stemphylium and Ulocladium species are best identified from dichloran chloramphenicol malt extract agar (DCMA; Andrews, 1992b) plates incubated for 7 days at 258C under lights. Conidial characteristics of Bipolaris species vary with type of medium. Species described here are best identified from tap water agar (TWA) containing a natural substrate such as sterilised wheat or millet seeds, or wheat straw. Drechslera species will grow well on DCMA but sporulation is poor. The best sporulation is achieved with V-8 juice (V-8 J) agar or TWA with one of the natural substrates mentioned above, incubated at 258C for 7–10 days under lights with a 12 h photoperiod. Light/dark periodicity is important as Drechslera cultures require light to produce conidiophores and darkness to produce conidia, possibly due to light inactivation of flavine necessary for conidial formation (Knan, 1971; Platt et al., 1977). For the identification of Trichoderma and Fusarium species, potato dextrose agar (PDA) is used. Fusarium species also require additional methods and media as outlined below.

4.8.5 Identification of Fusarium Species Fusarium isolates exhibit unusually high variability in colony morphology and also may deteriorate rapidly in culture. Thus, they should be identified as soon as possible after initial isolation with a minimum of subculturing to avoid deterioration. It is common practice to prepare cultures of Fusaria from single spores for growth on identification media, as this reduces both of these problems.

43

Single sporing. The technique for preparing single spore cultures is as follows (Nelson et al., 1983; Leslie and Summerell, 2006). Pour about 10 ml of 2% water agar into unscratched glass or plastic Petri dishes and allow to dry, either by holding the plates at room temperature for several days or by placing them inverted in an oven at 37–458C for about 30 min. Prepare a suspension of conidia in a 10 ml sterile water blank so that it contains 1–10 spores per low-power (10) microscope field when a drop from a 3 mm loop is examined on a slide. With experience, this concentration can be gauged simply by observing the turbidity of the suspension. Pour the suspension of spores onto a dried water agar plate, drain off the excess liquid and incubate in an inclined position at 20–258C for 18–20 h. After incubation, open the Petri dish, shake off any accumulated moisture droplets and examine under a stereomicroscope using transmitted light. The germinating conidia should be visible under 25 magnification. A dissecting needle with a flattened end and sharpened edges is used to cut out small squares of agar containing single, germinating conidia. These single conidia are then transferred on the agar blocks to the desired growth medium. If the original culture is contaminated with bacteria, a drop of 25% lactic acid may be added to the water blank. Allow this acidified spore suspension to stand for 10 min before pouring onto a water agar plate. Germination of acid-treated Fusarium conidia may be delayed by 24 h or more. Media. Two media have been used in this book for the identification of Fusarium isolates: potato dextrose agar (PDA) for colony characteristics and colour and dichloran chloramphenicol peptone agar (DCPA) for the development of diagnostic macro-, micro- and chlamydoconidia. A third medium, carnation leaf agar (CLA), is recommended by some Fusarium specialists for Fusarium cultivation and identification (Nelson et al., 1983; Leslie and Summerell, 2006). CLA is an excellent medium, on which most Fusarium species readily produce their diagnostic macroconidia. Production of macroconidia on DCPA is usually comparable with that on CLA, but microconidia and chlamydoconidia are often more plentiful on CLA due to greater production of aerial hyphae. DCPA is used in the present work rather than CLA,

44

however, because dried, gamma-irradiated carnation leaves are difficult to obtain in many localities. Inoculation and incubation. For identification by the methods used in this book, single spore cultures of Fusarium isolates should be prepared on agar blocks as outlined above, inoculated, one per plate, onto two plates each of PDA and DCPA and incubated at 258C for 7 days. Individual plates may be used for each medium, or alternatively divided plates may be used, with one medium on each half of the plate. Illumination during incubation is essential for the production of macroconidia. The light source may be diffuse daylight (not direct sunlight) or light from a bank of fluorescent tubes. A photoperiod of 12 h per day is normally used. Alternating temperatures of 20 and 258C have been recommended (Nelson et al., 1983; Leslie and Summerell, 2006) but are not essential. A simple light bank may be constructed from a standard 40 watt fluorescent fixture with two cool white tubes, suspended 0.5–l m above the laboratory bench or shelf supporting the cultures. The addition of a black light tube (e.g. Philips TL 40 W/80 RS F40BLB) is also desirable and in some cases essential to induce macroconidial or chlamydoconidial production.

4.8.6 Yeasts Yeast identification systems. Identification of foodborne yeasts remains a difficult task, as colony characteristics and microscopic morphology are of limited value. Generally it has been necessary to use biochemical and physiological tests such as fermentation of carbohydrates, assimilation patterns for a range of carbon and nitrogen sources and growth at various temperatures. Details of these methods and media may be found in Kurtzman and Fell (1998), Kurtzman et al. (2003) or Barnett et al. (2000). Molecular methods (see below) are being used increasingly in yeast identification, replacing time consuming biochemical testing. Identification using systems based on biochemical and physiological testing is complex and time consuming. However, a number of attempts have been made to assist those who wish to persevere with yeast identification. Several simplified systems

4 Methods for Isolation, Enumeration and Identification

have been published in the literature, and both automated and manual yeast identification systems are now commercially available. Dea´k and Beuchat (1987) published a simplified identification key (SIK) which included 215 species of foodborne yeasts. They subsequently modified their system, restricting it to the 76 species most frequently occurring in foods (SIM), and reported that it was much more successful than the API 20C system (BioMe´rieux, Marcy-l’Etoile, France) for identification (Dea´k, 1992). The SIM uses only two Petri dishes and three test tubes to examine each strain for ability to assimilate 10 carbon sources, fermentation of glucose, assimilation of nitrate and splitting of urea. These biochemical tests are supplemented by morphological observations. SIM separates the yeasts into six groups by a dichotomous key utilising the results of five key tests. Further tests are used to differentiate the yeasts in these six groups using secondary dichotomous keys. A dichotomous key to 25 common species of foodborne yeasts was published by Smith and Yarrow (1995), who used 17 biochemical and physiological tests to distinguish the species. Of the commercial systems available, the most widely used for foodborne yeasts are the Biolog, which is an automated system, and the BioMe´rieux ID32C yeast identification strips which can be read manually or automatically using the ATB system. BioMe´rieux also markets the fully automated VITEK 2 system, based on a card containing 64 tests. However, the database comprises only 46 clinically important species (Aubertine et al., 2006) and so has limited application to foodborne yeasts. The Biolog (Biolog Inc., Hayward, CA, USA; http://www.biolog.com/microID.html) utilises a yeast identification test panel (YT MicroPlateTM) consisting of a matrix of 8  12 wells. The first three rows contain 35 carbon source oxidation tests using tetrazolium violet as an indicator of oxidation. The next five rows contain carbon assimilation tests which are scored turbidimetrically against a negative control panel containing only water. The last row contains two carbon sources and tests for the co-utilisation of various carbon sources with D-xylose. The hardware (Biolog MicroStation Reader) consists of an automated plate reader coupled with a computer, which interprets the

4.9 Examination of Cultures

results and compares them with the resident database which currently includes 267 species. Manual interpretation of the Biolog plates is not recommended. This system has been designed with the food industry in mind, and the database contains all the common foodborne yeasts, unlike other systems which are usually aimed at the clinical market. The ATB ID32C system (BioMe´rieux, Marcyl’Etoile, France) is an automated system utilising the BioMe´rieux ID32C yeast identification strips. These strips contain 30 assimilation tests plus a positive (glucose) and a negative control well, all of which are inoculated with a yeast suspension of specified density. The strips are incubated at 308C for 48–72 h. As with the Biolog, the ATB automated system consists of a plate reader attached to a computer. The database associated with the ATB system contains 63 yeast species. The BioMe´rieux ID32C strips can be read manually, and the results enable identification of yeasts using published keys or computer identification programmes such as that of Barnett et al. (1996). This system may be used with reasonable success, particularly when the test results from the ID32C strips are supplemented with extra tests (glucose fermentation, urease production, nitrate utilisation, growth in 0.5 and 1% acetic acid and in 50 and 60% glucose, growth at 378C, production of pseudohyphae, ascospore formation and morphological observations) to give a more comprehensive base for the identification. Growth in 0.5 and 1% acetic acid indicates preservative resistance, and growth in 50 and 60% glucose gives an indication of ability to grow at reduced aw. Both these parameters are important in determining a yeast’s ability to cause spoilage in particular products. Identification using DNA sequencing is increasingly becoming the method of choice, as extensive databases such as GenBank are freely available for identification purposes. The 600– 650 nucleotide D1/D2 region of the large subunit (26S) ribosomal DNA is the most widely targeted section of the genome, and sometimes the ITS region may also be used (Kurtzman et al., 2003). Sequencing of the D1/D2 region, along with some supplementary physiological and biochemical tests, is the identification method currently used in our laboratory.

45

Even using the available systems, identification of yeasts still requires some specialist knowledge and interpretation and remains time consuming. Our experience indicates that no more than 12 species of spoilage yeasts are of real concern in foods. It is possible to differentiate these few species by relatively simple techniques, i.e. colony and microscopic morphology, growth on the standard media used for filamentous fungi, growth on other media which test for preservative resistance, ability to use nitrate as a nitrogen source and adaptation to high NaCl concentrations. Details of these techniques are given in Chapter 10.

4.9 Examination of Cultures As noted above, all cultures for identification should first be grown on the standard regimen described earlier. After 7 days incubation, the following examination should be carried out and then the general key to fungi in Chapter 5 should be used. That key will be of assistance even in the event of cultures failing to grow under one or more of the standard conditions.

4.9.1 Colony Diameters Measure the diameters of macroscopic colonies in millimetres from the reverse side (Fig. 4.4). Microscopic growth or germination at 58C is assessed by low power microscopy (60–100), by putting the 58C Petri dish on the microscope stage and examining by bright field transmitted light. Growth at 378C is assessed macroscopically only; germination of spores at 378C is an unreliable character.

4.9.2 Colony Characters Colony appearance can be judged by eye or with a hand lens, but examination is more effective if a stereomicroscope is used. Magnifications in the range of 5–25 are the most useful. Characters such as type and location of sporing structures and

46

4 Methods for Isolation, Enumeration and Identification

Fig. 4.4 Technique for measuring colony diameters by transmitted light

extent of sporulation are best gauged with the stereomicroscope. Reflected light is usually more effective than transmitted light. To determine colony colours, examine colonies by daylight or by daylight-type fluorescent light. In some genera, reference to a colour dictionary is helpful. The Methuen ‘‘Handbook of Colour’’ (Kornerup and Wanscher, 1978) has been used in this work and is highly recommended.

4.9.3 Preparation of Wet Mounts for Microscopy Fungi should always be examined microscopically as wet mounts rather than fixed and stained like bacteria. To prepare a wet mount, use an inoculating needle to cut out a small portion of the colony which includes sporing structures. Examination with the stereomicroscope can be an invaluable aid here. For freely sporing fungi with little mycelium, cut a piece of colony near the edge, where fruiting structures are young and spore numbers not excessive. Take structures which may enclose spores, i.e. cleistothecia, from near colony centres, where the probability of mature spores is highest. If the only differentiated parts of the colony appear to be

buried in the agar, e.g. pycnidia, take a sample of these with a small piece of the agar. Float the cut colony sample from the needle onto a slide with the aid of a drop of 70% ethanol. It may be necessary to tease out the specimen with the needle and the corner of a cover slip (square cover slips are best). Fungal specimens may be highly hydrophobic: the ethanol helps to wet the preparation, minimising the amount of entrapped air. When most of the ethanol has evaporated, add a drop of lactic acid (for phase or interference contrast optics) or lactofuchsin stain (see below) for bright field. Add a cover slip; if necessary remove excess liquid from the preparation by gently blotting with facial tissue or similar absorbent paper. The preparation is now ready for examination.

4.9.4 Staining A wide variety of stains are in use for mycological work. However, most are time consuming to prepare, or to use, or are slow to act, because fungal walls and spores are highly resistant to stains. By far the most effective stain for use in food mycology is lactofuchsin (Carmichael, 1955), which suffers from none of these faults. It consists of 0.1% acid fuchsin

4.9 Examination of Cultures

dissolved in lactic acid of 85% or higher purity. Young actively growing fungal structures are preferentially coloured bright pink, so sporing structures can usually be readily distinguished against a background of older mycelium. Cleistothecial initials, developing asci and maturing ascospores are also seen more readily in preparations stained with lactofuchsin. Like most other mycological stains, lactofuchsin is corrosive. Take care to clean it off microscope parts or skin! Be especially careful of the objective faces, because lactic acid will slowly corrode the relatively soft glass used in lenses.

4.9.5 Microscopes and Microscopy Choice of microscope. Fungal identifications inevitably involve microscopy. A high quality, binocular compound microscope is essential for serious mycological work. Bright field, phase contrast and interference contrast optics are all suitable. If bright field optics are used, preparations should be stained. Phase contrast optics avoid the need for staining, although as explained above, staining has merit for certain structures in any case. Resolution under phase contrast sometimes suffers from excessive halo effects because fungal structures are highly refractile. Experience suggests that bright field optics are more satisfactory than phase contrast for examining most fungi. Interference contrast is superior to both bright field and phase contrast and is strongly recommended. The very thin optical section cut by this system provides higher resolution of dense structures and the relatively low contrast is restful to use. The microscope should be equipped with at least four objectives: 6 for examining Petri dishes under low power (for germination, etc.); 10 or 16 for searching fields for sporing structures; 40 for examining structural details and measuring stipes, fruiting structures and large conidia; and 100 oil immersion for observation of details of spore attachment, surface texture, ornamentation of hyphae and spores and fine measurements. Oculars should be 10 or 12.5 and may with advantage be wide field and have a high eye point suitable for use with spectacles. One should be a focusing ocular,

47

equipped with an eyepiece micrometer, which is essential for measuring dimensions of spores and sporogenous structures. In the examination of fungal mounts, it is stressed that it is most important to use low power optics before succumbing to the temptation to use oil immersion. The principal reason is that fungal preparations usually remain as small clumps and do not disperse as bacteria do. Only under low power is the search for the optimal area of the slide for the observation of fruiting structures likely to be rewarded. Once a suitable area is located under the 10 or 16 objective, move to the 40. This should be the lens most used; microscope optics are such that only the finest details of ornamentation can be observed more effectively under oil immersion than at this magnification. Aligning the microscope. Correct alignment of the microscope is essential, so that its resolution is as high as possible and it can be used for long periods without discomfort. An incorrectly aligned microscope will lead to poor observation, discomfort, fatigue, headache and eye strain. A person of normal visual acuity should be able to use a correctly aligned instrument throughout a whole working day without discomfort. The steps to correctly align a microscope are given below. They should be read in conjunction with the microscope manufacturer’s instructions. 1. Mount a slide on the stage and bring it into approximate focus. If a prepared slide is not available, a slide marked with a marking pen or ink line is a satisfactory substitute. 2. Close the microscope’s field diaphragm (the one at the microscope base nearer the light source). The image of the diaphragm opening should now be visible in the microscope field. If it is, first focus it with the condenser focusing knob and then centre it in the field with the condenser centring screws. If the diaphragm opening cannot be seen, first rack the condenser up and down and watch to see if the opening becomes visible; if it does not, rack the condenser to its highest position and then slowly open the field diaphragm until the opening comes into view. Centre the diaphragm approximately and proceed as above. 3. For bright field optics, the condenser diaphragm should be adjusted each time the objective power is changed. Remove one ocular; close the

48

condenser diaphragm so that the field seen down the open tube is about two-thirds its maximum size. With phase contrast and interference contrast systems, this adjustment is less critical. The preceding steps align the microscope itself and should be checked frequently. If optimal illumination is desired, each step should be carried out for each new slide and each objective change. As a routine habit, the whole process should take only a few seconds. If the available microscope is not equipped with a built-in light source, a field diaphragm and a fully centring condenser, it is unlikely to be satisfactory for the identification of small spored fungi such as Penicillium species. The following steps are designed to align the observer with the microscope, compensating for individual differences in sight. Provided settings on the microscope are remembered, these steps need be carried out only occasionally, to check that visual acuity has not altered. Different settings will be needed for an individual with and without spectacles or contact lenses. 4. Assuming the microscope is binocular, pull the oculars out to their greatest distance apart and then, while watching a focused field, move them gradually together until a single circular field is seen without strain or head movement. Note the distance on the scale between the oculars; this is the individual’s interpupillary distance. Repeat this operation two or three times until satisfied that the correct distance has been found. This distance should always remain the same and be similar on any microscope. 5. Under the 40 or 100 objective, locate a tiny, readily recognised point on the slide and focus on it. Take a piece of white card and place it between the focusing ocular and the corresponding eye. Leave the eye open. Now focus the tiny point with the other eye, carefully, with the microscope fine focus. Next, transfer the white card to the other ocular and, using the focusing collar beneath the ocular, refocus the tiny point. Remove the card and note the setting on the scale. Repeat until satisfied the correct setting has been found. 6. On some microscopes, the eyepiece micrometer can be focused independently. Use the focusing system on the ocular itself to focus the micrometer. Note the setting on the scale on the side of the ocular.

4 Methods for Isolation, Enumeration and Identification

Always check the settings on the microscope before use and after making measurements with the micrometer. It is very easy to upset the ocular alignment when measuring.

4.10 Preservation of Fungi Many fungi are stable in culture and can be subcultured repeatedly without apparent change or deterioration. Others, especially Fusarium species and other plant pathogens and some mycotoxigenic species, will degenerate rapidly after only a few transfers. For stable isolates that are used routinely in the laboratory, storage on agar slopes is satisfactory. Many freely sporing fungi will survive for several months, and sometimes much longer, when stored at 1–48C on a medium such as CYA. One of the hazards of storage at these temperatures is contamination by psychrophilic Penicillium or Cladosporium species. Storage at freezer temperatures (–18 to –208C) prevents such contamination, but some fungi do not survive well under these conditions. Storage at room temperatures, at 108C or above, for long periods is not advisable because of the likelihood of invasion by culture mites (see below).

4.10.1 Lyophilisation For unstable cultures, and indeed for the long term storage of any food spoilage fungi, freeze drying or lyophilisation is probably the best method of preservation. Many commercial systems are now available for carrying out this process. A satisfactory menstruum for lyophilisation of most fungi is l.5 normal strength reconstituted nonfat milk powder (15% in distilled water). For fungi with hydrophobic conidia, such as Aspergillus and Penicillium, a small amount of detergent (0.05%) such as polysorbitan 80 (Tween 80) should be added to the milk. Dispense the milk in 10 ml lots in small tubes or 12.5 ml (0.5 oz) McCartney bottles, and sterilise by steaming for 20 min on three successive days (the Tyndallisation process). The milk must be stored at 208C or above between steamings, to permit bacterial spores to germinate.

4.10 Preservation of Fungi

Occasionally bacterial spores will survive this process: it is advisable to store the milk at room temperature for some days after Tyndallisation. Any bottles which show clotting or other breakdown should be autoclaved and discarded. Autoclaving at 1218C, even for 10 min or less, is not recommended, as browning will occur and browning compounds are known to be inhibitory to microorganisms. Most common spoilage fungi survive lyophilisation well. However, in our experience and that of others (Mikata et al., 1983; Smith and Onions, 1994), some yeasts, plant pathogens and xerophiles do not. Storage of lyophilised ampoules at refrigeration temperatures (0–48C) is recommended, but room temperature is probably satisfactory provided sunlight is avoided and temperatures do not exceed 308C. Some laboratories routinely store lyophilised cultures at –18 to –208C. Strains being maintained for a particular trait or utilised for a specific purpose such as system testing or metabolite production should always be lyophilised. Continued subculturing often leads to deterioration or loss of the desired character. The ability of a strain to produce a particular mycotoxin, for example, may decrease with each transfer. Isolates should be lyophilised as soon as possible after primary isolation to prevent degeneration.

4.10.2 Other Storage Techniques A variety of systems other than lyophilisation have been proposed for long term storage of fungi. Of these, liquid nitrogen storage has found most acceptance with major culture collections. This type of storage appears to be superior to any other for plant pathogens and fungi which will not sporulate in pure culture. However, liquid nitrogen systems are expensive to establish and maintain and are only suitable for large collections. Freezer units which run at very low temperatures (–808C or below) are available and are well suited to the needs of culture collections. In our laboratory we routinely store cultures at –808C, using glycerol (60–80%) as a cryoprotectant. Spore suspensions are prepared by taking conidia or ascospores from a freely sporulating sector of the colony, dispersing them in the glycerol then freezing in screw capped

49

cryovials. An 80% solution of glycerol remains viscous at –808C, which enables cultures to be removed from the freezer for subculture without the need for defrosting. For smaller laboratories that do not have access to lyophilisation, liquid nitrogen or ultra-low temperature storage, some simple techniques exist that can be used to maintain fungal cultures over relatively long periods (e.g. 1–10 years) without the need for subculturing. Water storage. A simple and inexpensive method of fungal culture preservation is storage of agar blocks in water (Smith and Onions, 1994). Small agar blocks (7–10 mm2) are cut from the growing margin of a young fungal colony and placed in sterile water in a bottle such as a Bijou bottle (6.25 ml or 0.25 oz McCartney bottle). The rubber lined cap is screwed down and the bottles stored in a cool room (1–108C). Cultures may be revived by removal of a block and placing it on a suitable growth medium. Using this method, cultures have been reported to remain viable and retain their characteristics for up to 7 years (Boeswinkel, 1976; Smith and Onions, 1994). Some yeasts may be maintained by storing as suspensions in water (Kirsop, 1984). Growth from a late logarithmic slant culture is suspended in sterile distilled water and transferred to a sterile container so that 90% of the volume is filled with the suspension. Containers are stored at room temperature. Survival of some Candida, Saccharomyces, Cryptococcus, Rhodotorula and Schizosaccharomyces species for up to 4 years has been reported (Kirsop, 1984). Silica gel storage. Many fungal cultures may be maintained for long periods (often more than 10 years) by drying spore suspensions onto silica gel (Smith and Onions, 1983; Smith, 1984). This method is not suitable for mycelial cultures, but can be used with some success for yeasts (Kirsop, 1984). As silica gel liberates heat when moistened, the technique depends on keeping the cultures cool enough to avoid damaging the spores during preparation. Medium grain plain (non-indicating) silica gel of 6–22 mesh is placed in suitable glass bottles (Bijou or McCartney bottles) to a depth of about 1 cm and sterilised, either by dry heat at 1808C for 2–3 h or by autoclaving at 1218C for 15 min. Autoclaved silica gel must be thoroughly

50

dried in an oven before use. Bottles are precooled by placing in a tray of ice or refrigerating for 24 h, then transferring to an ice tray for inoculation. Suspensions of fungal spores or yeasts are prepared in sterile skimmed milk (as for lyophilisation, above) and the suspension added to the cooled silica gel to wet three-quarters of it. The bottles and gel are allowed to dry at room temperature for 10–14 days with caps slightly loosened. Caps are then screwed down and the bottles stored at 48C (storage at room temperature is also satisfactory) in air-tight containers over indicating silica gel to absorb any moisture. Cultures are revived by shaking a few crystals of silica gel onto a suitable growth medium (broth culture may be better for yeasts). Survival varies according to species or even strain, but survival of yeasts for up to 5 years (Kirsop, 1984) and fungi for more than 10 years (Smith and Onions, 1983) has been reported.

4.11 Housekeeping in the Mycological Laboratory Like any other microbiological laboratory, a mycological laboratory should be kept in a clean condition. Discard unwanted cultures regularly and dispose of them by steaming or autoclaving. Wipe bench tops regularly with ethanol (70–95%). Floors should be wet mopped or polished only with machines equipped with efficient vacuum cleaners and dust filters. Where possible store food and plant materials away from the laboratory. Open Petri dishes carefully. Use small inocula on wet needles. Transport Petri dishes to the stereomicroscope stage before removing lids. Do not bump cultures during transport. Contrary to popular belief, a well-run mycological laboratory is not a source of contamination to bacteriological laboratories. The air in a mycological laboratory should not carry a significant population of fungal spores. The reverse problem can occur, however, because bacteria multiply more rapidly than do fungi. Bacterial spores are often present in food laboratories, readily infect fungal plates and can rapidly outgrow and inhibit fungal mycelia, especially at 378C.

4 Methods for Isolation, Enumeration and Identification

If for any reason fungal spore concentrations do build up in a laboratory and cause an unacceptable level of contamination, the air should be purified. The simplest technique is to spray the air throughout the laboratory with an aerosol before it is closed in the evening. Any aerosol spray, such as a room deodoriser or air freshener, is effective. Aerosol droplets entrain fungal spores very efficiently and carry them to the floor. A more drastic and effective treatment in cases of severe contamination is to spray a solution of 2% thymol in ethanol around the room and close it for a weekend. The spray is rather pungent, and while not harmful to humans, it effectively kills fungal spores and mites (see below). Do not leave cultures on benches before fumigation.

4.11.1 Culture Mites A major hazard in growing and maintaining fungal cultures is the culture mite. Many species of mites live on fungal hyphae as their main or sole diet in nature and find culture collections an idyllic environment. Mites crawl from culture to culture, contaminating them with fungi and bacteria as they go or, given long enough, may eat them out entirely. Mites are very small (0.05–0.15 mm long), usually just visible to the observant naked eye. They are arachnoids, related to spiders, and hermaphroditic. Each mite leaves a trail of eggs about half adult size as it goes. Eggs hatch within 24 h and reach adulthood within 2 or 3 days. The damage an unchecked mite plague can do to a fungal culture collection has to be experienced to be believed. The most common sources of mites are plant material, soil, contaminated fungal cultures and mouldy foodstuffs. Mites can also be carried on large dust particles. Building work near a laboratory almost always induces a mite infestation. The avoidance of losses due to mites requires constant vigilance. Always watch for telltale signs, such as contaminants growing around the edges of a Petri dish, a ‘‘moth-eaten’’ appearance to colonies or ‘‘tracks’’ of bacterial colonies across agar. Examination of suspect material or cultures under the stereomicroscope will readily reveal the presence of mites and mite eggs.

4.11 Housekeeping in the Mycological Laboratory

Adult mites are rapidly killed by freezing, and mite eggs will only survive 48–72 h at –208C. Cultures contaminated by mites can often be recovered by freezing for 48 h, then subculturing from uninfected portions of the culture with the aid of the stereomicroscope. Suspect food and other samples being brought into the laboratory should also be frozen for 24 h to destroy mites before enumeration or subculturing is carried out. Infestation by mites can be minimised by good housekeeping, i.e. by avoiding accumulation of dust or old cultures in the laboratory. It is also good practice to handle and store food and plant samples well away from areas where fungi are inoculated and incubated. To control a mite infestation, remove all contaminated material, including cultures. Freeze Petri dishes and culture tubes which must be recovered; autoclave, steam or add alcohol to all others. Clean benches thoroughly with sodium hypochlorite (household bleach) or 70% ethanol. Incubators can be disinfested with aerosol insecticides.

4.11.2 Problem Fungi There are three fungal invaders which should be watched for carefully in a food mycology laboratory: Aspergillus fumigatus, Rhizopus stolonifer and Chrysonilia sitophila. The first is a human pathogen; the others can cause a contamination chain which is difficult to break. Aspergillus fumigatus may cause invasive aspergillosis in the lungs or serious allergenic responses in some individuals. It is sound practice to immediately kill and discard cultures of this fungus as soon as it is recognised. On no account should it be used for experimental studies in food spoilage or biodeterioration without precautions to prevent dissemination of spores. The morphology of A. fumigatus is described in detail in Chapter 8, but it is readily recognisable in the unopened Petri dish:

 colonies are low, dull blue and broadly spreading, with a velvety texture;

 growth is very rapid at 378C, covering a Petri dish in 2 days;

 long columns of blue conidia are readily seen under the stereomicroscope.

51

Rhizopus stolonifer is a ubiquitous fungus in many kinds of foods. It grows rampantly at 258C, filling a Petri dish with sparse, dark mycelium in 2 days. It produces barely macroscopic aerial fruiting structures which are at first white, then become black. Given seven undisturbed days, it sheds dry, black spores outside the Petri dish, providing an effective inoculum for a continuous chain of future contamination. Once such an infection occurs, it is essential to carefully place the contaminated plate in a suitable container such as a plastic bag before transporting it to steamer or autoclave. Then use 70% ethanol to clean areas on which the plate had been incubated or placed. At daily intervals, carefully examine all plates inoculated subsequently, discarding any which show Rhizopus contamination, until infection ceases. Spraying the air with aerosols or thymol in ethanol will assist. Unlike A. fumigatus, R. stolonifer is not pathogenic. The third problem fungus, Chrysanilia sitophila, is known as ‘‘the red bread mould’’. It used to be common, but due to changes in manufacturing practice, it is now encountered less frequently, either in the bakery or in the laboratory. Like Rhizopus stolonifer, it grows with great rapidity at 258C. It forms a thin, pink mycelial growth across a Petri dish, clearly following the oxygen gradient which leads to the open air. It will force its way unerringly between dish and lid and, once outside, will produce masses of pink spores, which are quickly shed and build up around the base of the Petri dish. Decontamination relies on the same techniques as for Rhizopus: C. sitophila is probably the more difficult fungus to eradicate. Again, apart from its nuisance value in the laboratory, it is harmless.

4.11.3 Pathogens and Laboratory Safety While it must be said that any fungus which is capable of growth at 378C is a potential mammalian pathogen, the physiology of the healthy human is highly resistant to nearly all of the fungi encountered in the food laboratory. Nevertheless, fungi which can grow at 378C should be treated with caution. In particular, the habit of sniffing cultures is to be avoided wherever possible. It is true that odours produced by fungi have been used quite frequently as taxonomic criteria, especially in older

52

publications, but their subjective and ephemeral nature makes them of little value for this purpose, and the risks involved are serious. Some laboratories regard fungal volatiles as such a serious risk that cultures such as Penicillium and Aspergillus species are handled in a biohazard cabinet. Many types of fungal spores are allergenic or carry mycotoxins. Inhalation of fungal spores should be avoided as far as possible. Of the fungi described in this book, only the Aspergilli normally pose any direct threat to health. Aspergillus fumigatus has already been mentioned, and care should also be taken when handling cultures of Neosartorya species, which are closely related to

4 Methods for Isolation, Enumeration and Identification

A. fumigatus and also grow prolifically at 378C. Other Aspergillus species, particularly A. flavus, A. niger and A. terreus, have been isolated from human specimens from time to time. For more details, see de Hoog et al. (2000). These species appear to be mainly opportunists and pose little threat to healthy people. Careful handling and good housekeeping are all that are required. However, immunocompromised individuals are in a different category. It is increasingly evident that such people have little resistance to fungal infection. Persons suspected to be immunocompromised, regardless of the cause, should not work in a mycology laboratory nor indeed be permitted knowingly to enter one.

Chapter 5

Primary Keys and Miscellaneous Fungi

Principles underlying fungal classification have been outlined in Chapter 3, including a brief overview of the relevant divisions of the Kingdom Fungi and their principal methods of reproduction. Some further detailed information is necessary in this chapter to assist in the use of the keys which follow. Ascomycetes. As discussed in Chapter 3, Ascomycetes produce ascospores in asci (Fig. 3.2). One genus, Byssochlamys, produces asci which are unenclosed; all other genera of relevance here produce asci in some kind of fruiting body, or ascocarp. The two kinds of ascocarp commonly seen in food spoilage fungi, the cleistothecium and the gymnothecium, have been described and illustrated in Chapter 3 (Fig. 3.3). Both types of ascocarp are usually pale or brightly coloured, not dark, and release ascospores by rupturing irregularly. Of genera relevant here, cleistothecia are produced by Emericella, Eurotium, Eupenicillium, Monascus and Neosartorya, and gymnothecia by Talaromyces. A third class of ascocarp, less commonly encountered in foodborne fungi, is the perithecium. Perithecia have cellular walls like cleistothecia, but are distinguished by the presence of an apical pore or ostiole through which asci or ascospores are liberated; also asci are long and clavate with ascospores arranged linearly within them. In the one perithecial genus of interest here, Chaetomium, the perithecia are black and have stout hyphae attached to the walls (Fig. 5.10). Conidial fungi. The strictly conidial fungi, also known as Fungi imperfecti or Deuteromycetes or anamorphic fungi, possess an amazing variety of ways of producing conidia. Terminology for structures bearing conidia and for conidia themselves has

become astonishingly complex in recent years; fortunately most of it is not essential for the recognition of the genera discussed in this text. Terms which are important in the keys which follow are described below. A fundamental division within the asexual fungi separates genera which form conidia aerially, grouped in the class Hyphomycetes, from those in which conidia are borne in some sort of enveloping body, the class Coelomycetes. Hyphomycetes. Fungi have developed seemingly endless ways of extruding or cutting off conidia, solitarily or in chains, from fertile cells which themselves may be borne solitarily or aggregated into more or less ordered structures. Hyphomycete taxonomy attempts to thread a way through this maze. In general, type and degree of aggregation of the fertile cells, and type of conidium, provides the basis for generic classification, while details of these characters and of spore size, shape and ornamentation are more commonly used to distinguish species. Features of conidia used in the keys in this work are length, septation, ornamentation and colour, particularly whether walls are light or dark. The method of conidium formation (ontogeny) is seldom emphasised here, because terminology is complex and distinctions may not be obvious. The principal point to note is the disposition of conidia: they may be borne solitarily, i.e. just one conidium per point of production; singly, i.e. successively from a single point, but unattached to each other; or in chains. Solitary conidia are borne on a relatively broad base and usually adhere to the fertile cell. Conidia formed in chains are usually extruded from a small cell of determinate length, often a

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_5, Ó Springer ScienceþBusiness Media, LLC 2009

53

54

phialide, which in most genera narrows to a distinct neck. Conidia borne singly may be extruded in this same manner, or be borne by extrusion from a pore in a hypha or fertile cell, or be cut off by hyphal fragmentation. Phialidic Hyphomycetes. Hyphomycetes may produce phialides solitarily (the genus Acremonium) or in loosely ordered structures (Trichoderma) or highly ordered structures (Aspergillus, Penicillium and related genera). Genera of interest here with less ordered phialidic structures can mostly be differentiated by macroscopic characters, e.g. colony diameters and colours. However, differentiating genera with highly ordered phialidic structures will necessitate careful microscopic examination. Phialides in Aspergillus, Penicillium and related genera are sometimes borne directly on a stalk or stipe which arises from a hypha; in other species, however, the phialides are borne from supporting cells, termed metulae (sing. metula) and in some species the metulae may in turn be supported by other cells, termed branches or rami (sing. ramus). The whole structure, including the stipe, is called a conidiophore. In Aspergillus, stipes are always robust, with thick walls and usually without septa; the stipe terminates in a more or less spherical swelling, the vesicle, which bears phialides, or metulae and phialides, over most of its surface. In Aspergillus, phialides (and metulae) are always produced simultaneously, and this feature can readily be recognised by examining young developing conidiophores (Fig. 8.1a). Similar structures, though smaller, are produced by some Penicillium species: these are clearly distinguished from Aspergillus species by stipes which are septate and by phialides which are produced over a period of time (successively; Fig. 8.2b). Most Penicillium species, and those of related genera, do not produce phialides on vesicles, but in a cluster directly on a stipe, or on metulae and/or rami. The fruiting structure in Penicillium and related genera is termed a penicillus, while that in Aspergillus (for want of a better term) is called a head (or more recently, an aspergillum, Klich, 2002). Coelomycetes. As noted earlier, Coelomycetes produce conidia within an enveloping body, termed a conidioma (pl. conidiomata). In Petri dish culture, conidiomata are produced on or just under the agar surface and are macroscopically visible, usually being 100–500 mm in diameter. Two kinds of conidioma are important here: the pycnidium, a more

5 Primary Keys and Miscellaneous Fungi

or less spherical body with one or more pores (ostioles) through which conidia are released, and the acervulus, a flat body from which conidia are released by lifting or rupturing of a lid. The majority of Coelomycetes are pathogens on plants and many have not been studied in pure culture. In consequence, their taxonomy is difficult and genera and species are often poorly delimited. For a complete account of Coelomycete taxonomy see Sutton (1980). Yeasts. Yeasts are fungi which have developed the ability to reproduce by forming single vegetative cells by budding or, in a few species, by fission, in a manner similar to bacteria. Like bacteria, and unlike fungal spores, such cells are metabolically active and may in turn reproduce by budding (or fission). Yeast cells may survive for long periods both in culture and in nature. In consequence many yeasts produce true spores rarely or not at all. Yeasts are readily distinguished from filamentous fungi on the agar plate by their soft-textured colonies and limited growth. They are usually also readily distinguished from bacteria by their raised and often hemispherical colonies, white or pink colours and lack of ‘‘bacterial’’ odour. If in doubt, make a simple wet mount of a colony in water or lactofuchsin, add a cover slip and examine with the oil immersion lens. Yeast cells are larger than bacteria, measuring at least 3  2 mm and are nonuniform in size. If the culture is not too old, some cells will usually show developing buds. Yeasts cannot be classified solely by morphological features or growth on the standard media, and so are considered in a separate chapter (Chapter 10).

5.1 The General Key The taxonomic terms discussed above will enable the use of the general and miscellaneous keys which follow, although some other taxonomic terms may be introduced in discussions of particular genera. It is emphasised that these keys are designed for use on isolates which have been incubated for 7 days on the standard plating regimen outlined in Chapter 4. Colony diameters should be measured in millimetres from the reverse side by transmitted light. The general key has been designed to be as simple as possible and suitable for routine use, but it should be read in conjunction with the notes below it.

5.1 The General Key

55

General key to food spoilage fungi 1

No growth on any standard medium in 7 days Growth on one or more standard media

Chapter 9 – ‘‘Xerophilic fungi’’ 2

2 (1)

Colonies yeasts, either recognisably so on isolation or in culture, i.e. colonies soft, not exceeding 10 mm diam on any standard medium Growth filamentous, exceeding 10 mm diam on one or more standard media

Chapter 10 – ‘‘Yeasts’’ 3

3 (2)

Growth on CYA and/or MEA faster than on G25N Growth on G25N faster than on CYA and MEA

4 (3)

Hyphae frequently and conspicuously septate Hyphae lacking septa or septa rare

5 (4)

No mature spores present in 7 days Mature spores present in 7 days

6 9

6 (5)

Immature fruiting structures of some kind present No fruiting structures (or spores) detectable by lowpower microscopy or wet mounts from CYA or MEA

7

5 Chapter 6 – ‘‘Zygomycetes’’

7 (6)

Colonies and fruiting structures white or brightly coloured Colonies or fruiting structures dark

8 (7)

Colonies and fruiting structures white Colonies or fruiting structures brightly coloured

9 (5)

Spores (conidia) less than 10 mm long, borne in chains on clustered fertile cells (phialides), on well-defined stipes Spores (conidia) of various sizes, borne singly or solitarily, or if borne in chains, then chains not in aggregates

10 (9)

11 (10)

Conidia blue or green, phialides produced successively on vesicles, vesicles less than 10 mm diam, stipes usually septate Conidia variously coloured, phialides and/or metulae produced simultaneously on vesicles, vesicles larger than 10 mm diam, stipes nonseptate

Couplet 1. No growth on any standard medium indicates an extreme xerophile, i.e. Xeromyces bisporus or a Chrysosporium species, or a nonviable culture. Next inoculate culture onto MY50G for 7 days at 258C. If growth occurs, enter the key in Chapter 9, ‘‘Xerophilic Fungi’’; no growth on MY50G indicates a nonviable culture. Chrysosporium and Xeromyces

See section on ‘‘Miscellaneous fungi’’ below 8 Continue incubation; when spores mature, refer to section on ‘‘Miscellaneous fungi’’ below Chapter 8 – ‘‘Aspergillus and its teleomorphs’’ Chapter 7 – ‘‘Penicillium and related genera’’

Phialides or metulae and phialides borne on more or less spherical swellings on the stipe apices Phialides borne on penicilli, i.e. on unswollen stipes with or without intervening metulae and rami

5.1.1 Notes on the General Key

4 Chapter 9 – ‘‘Xerophilic fungi’’

10 See section on ‘‘Miscellaneous fungi’’ below

11 Chapter 7 – ‘‘Penicillium and related genera’’

Chapter 7 – ‘‘Penicillium and related genera’’

Chapter 8 – ‘‘Aspergillus and its teleomorphs’’

isolates are usually white or rarely golden brown. If the original culture used as inoculum is coloured other than pure white or golden brown, it is probably nonviable. Couplet 2. Yeasts are usually readily distinguished by slow growth, soft, easily sampled colonies, small spherical to ellipsoidal cells, often of variable size and shape and by reproduction by budding. See Chapter 10, ‘‘Yeasts’’ for identification procedures.

56

5 Primary Keys and Miscellaneous Fungi

Couplet 3. The ability to grow more rapidly on G25N than on CYA or MEA indicates a xerophile (at least for keying purposes here). Check the key in Chapter 9, ‘‘Xerophilic Fungi’’. Some isolates can be identified from the standard plates, while others will require growth on CY20S or MY50G for identification. Couplet 4. The absence of septa in young, growing hyphae indicates an isolate belongs to subkingdom Zygomycotina, discussed here in Chapter 6 ‘‘Zygomycetes’’. Couplets 5, 6. Some isolates from a variety of fungal genera will not produce spores on the standard media in 7 days. Continue to incubate such cultures, preferably in diffuse daylight such as a laboratory window sill, at temperatures near 258C. Also inoculate such cultures onto two or three plates of DCMA and incubate these at 258C or thereabouts in darkness and in diffuse daylight or, if possible, under fluorescent illumination (see Chapter 4). After 1–2 weeks, check again for spores or fruiting bodies. If such structures are not seen, the isolate is unlikely to be significant in foods. Apparently asporogenous cultures should also be checked with a stereomicroscope while scraping up a sector of the colony with a needle. Fruiting bodies submerged in the agar will sometimes become visible with this technique. Couplet 7. Some isolates which produce white or brightly coloured fruiting bodies also produce very sparse aerial conidial structures which are easily overlooked. Check such cultures carefully with the stereomicroscope; if conidial structures are found, make a wet mount and reenter the key at Couplet 5.

A finely drawn glass needle will sometimes be of assistance in making mounts from delicate conidial structures on the colony. Nearly all dark fruiting structures encountered will mature at 258C within 2 weeks. Light does not usually influence this process. When mature spores are formed, refer to the following section.

5.2 Miscellaneous Fungi In this section are considered the genera which do not logically fit into some larger grouping considered elsewhere. Some are important in specific food spoilage problems, others are found in particular habitats such as cereals, while still others represent the aerially dispersed fungal flora found as ubiquitous contaminants or saprophytes. As will be seen, they are a very heterogeneous collection. Most fungi significant in food spoilage or food contamination and not treated in other chapters are included here. It is inevitable, though, that occasional isolates from foods will not belong to the genera considered in this section. The key has not been designed to take account of this, as it would be a practical impossibility. So when an isolate appears to key out satisfactorily, it must be checked against the description to confirm the identification. Some isolates will of course belong to a recognisable genus, but not the species described; in that case the references indicated will provide further information. The miscellaneous fungal genera are considered in alphabetical order following the key.

Key to miscellaneous genera 1

Colonies on CYA and MEA not exceeding 60 mm diam in 7 days Colonies on CYA or MEA exceeding 60 mm diam in 7 days

2 12

2 (1)

Conidia borne within a fruiting body on or beneath the agar surface Conidia borne from aerial or surface hyphae

3 (2)

Mycelium and conidia hyaline or brightly coloured Mycelium and/or conidia dark coloured

4 11

4 (3)

Conidia with a single lateral septum Conidia nonseptate or with more than one septum

Trichothecium 5

5 (4)

Conidia borne from gradually tapering fertile cells (phialides) Conidia borne directly on hyphae or by budding or hyphal fragmentation

Phoma 3

6 7

5.2 Miscellaneous Fungi

6 (5)

Colonies exceeding 50 mm diam on CYA Colonies not exceeding 50 mm diam on CYA

7 (5)

Colonies exceeding 45 mm diam on MEA; conidia borne solely by the breakup of hyphae to form arthroconidia Colonies not exceeding 40 mm diam on MEA; conidia not exclusively arthroconidia

8 (7)

9 (8)

Conidia exceeding 12 mm long; developing cleistothecia, fist-like on arm-like stalks, also present Conidia not exceeding 12 mm long; no evidence of cleistothecial development Conidia yeast-like, borne on spicules (small projections) from hyphae, or by budding; conidia 5 mm or less long Some conidia yeast-like, borne by budding; hyphal fragments, arthroconidia and/or chlamydoconidia also present; conidia >5 mm long

10 (9)

Budding hyphal fragments (10–50 mm long) present Hyphal fragments absent; or if present, not budding

11 (3)

Colonies low, mucoid and yeast-like, becoming grey to black in both obverse and reverse Colonies dry and velutinous, obverse green, reverse olive or deep blue black

57

Fusarium Acremonium

Geotrichum 8

Monascus 9

Endomyces 10 Hyphopichia Moniliella

Aureobasidium Cladosporium

12 (1)

Spores borne within an enclosed fruiting body on or under the agar surface Spores borne from aerial or surface hyphae

13 17

13 (12)

Spores consistently less than 15 mm long The larger or all spores more than 15 mm long

14 15

14 (13)

Fruiting bodies perithecia with stout, black hyphae attached to the walls Fruiting bodies pycnidia, without attached hyphae

15 (13)

Fruiting bodies roughly spherical (pycnidia) Fruiting bodies flat (acervuli)

16 (15)

Conidia hyaline or brightly coloured, nonseptate, without terminal appendages Conidia dark, with three or four septa and with spike-like, sometimes branched, terminal appendages

Chaetomium Phoma Lasiodiplodia 16 Colletotrichum Pestalotiopsis

17 (12)

Colonies and conidia hyaline or brightly coloured Colonies and/or conidia dark coloured

18 22

18 (17)

Colonies with grey or green areas Colonies white, orange, pink or purple

19 20

19 (18)

Colonies green Colonies grey

20 (18)

Colonies low and persistently white Colonies floccose, white or becoming brightly coloured

Geotrichum 21

21 (20)

Colonies predominantly orange, orange conidia shed profusely around the Petri dish rim Colonies white, pink or purple, sporulation on MEA weak or absent, better on DCPA under lights

Chrysonilia

22 (17)

Conidia consistently less than 15 mm long Conidia frequently exceeding 15 mm long

Trichoderma Botrytis

Fusarium 23 25

58

23 (22)

5 Primary Keys and Miscellaneous Fungi

Conidiophores long, branched, apically swollen, bearing closely packed pale brown conidia Conidiophores short or ill-defined, dark brown or black conidia borne irregularly

Botrytis 24

24 (23)

Conidia dark brown, often with a lighter coloured band around the periphery Conidia uniformly jet black

Arthrinium Nigrospora

25 (22)

Conidia approximately spherical Conidia elongate

Epicoccum 26

26 (25)

Conidia with transverse septa (or thick walls between cells) only Larger conidia with both transverse and longitudinal septa or irregularly septate

27 30

27 (26)

Conidia clavate (club shaped), often with long hyphal appendages at the apices; found almost exclusively on rice Conidia cylindroidal, ellipsoidal or an elongate ‘‘D’’ shape; source more general

Trichoconiella 28

28 (27)

Conidia cylindroidal, with parallel sides except at the terminal cells Conidia fusoid, narrowing from the central cells to the terminal cells, often bent or ‘‘D’’ shaped

Drechslera

29 (28)

Conidia not exceeding 40 mm long Conidia usually exceeding 40 mm long

Curvularia Bipolaris

30 (26)

Conidia clavate (club shaped) Conidia spherical to roughly ellipsoidal or short cylindrical

Alternaria 31

31 (30)

Conidia often tapering towards the base and sometimes pointed, i.e. pyriform or apiculate Conidia spherical to short cylindrical, not tapering from base to apex, with rounded ends

5.3 Genus Acremonium Link Commonly referred to as Cephalosporium Corda in pre-1970 literature, Acremonium is a large and varied genus characterised by the production of small, hyaline, single-celled conidia borne singly, i.e. successively but not connected to each other, from solitary phialides. A variety of species have been recorded from foods from time to time. One species, A. strictum, described here, is of relatively common occurrence. In this species, the phialides gradually taper to the apices without basal thickening or formation of a distinct neck, and conidia aggregate in balls of slime. Under the stereomicroscope, the slime balls look like large single spores, but their true nature becomes evident in wet mounts. A. strictum appears mainly in the earlier food literature under the name C. acremonium. As noted by Domsch et al. (1980), this name has been used for a variety of species, so that reports on physiology and occurrence are unreliable (Fig. 5.1).

Acremonium strictum W. Gams

29

Ulocladium Stemphylium

Fig. 5.1

Cephalosporium acremonium (name of uncertain application; no valid authority).

Colonies on CYA 20–30 mm diam, white or orange to pink, dense to floccose or funiculose; reverse pale or with orange to pink tones. Colonies on MEA 13–20 mm diam, similar to those on CYA or of slimy texture. Colonies on G25N <5 mm diam, usually 1–2 mm, of white mycelium. Sometimes growth at 58C. No growth at 378C. Conidia borne successively but singly from the apices of long, solitary, usually unbranched phialides, aggregating in a slime ball at the phialide tip, cylindrical to ellipsoidal, hyaline, 3–6  1–2 mm, smooth walled. Distinctive features. See genus preamble. Taxonomy. As noted above, Acremonium strictum is the correct name for the fungus commonly called C. acremonium in the earlier food literature (Gams, 1971).

5.3 Genus Acremonium

59

Fig. 5.1 Acremonium strictum (a) colonies on CYA and MEA, 7 d, 258C ; (b, c) phialides, bar = 10 mm; (d) conidia, bar = 5 mm

Physiology. Mycelial growth and sporulation of Acremonium strictum were found to be enhanced by the addition of mannitol to MEA (Teixeira et al., 2005). Mycotoxins. A. strictum and other Acremonium species encountered in foods are not known to produce mycotoxins. Ecology. The main source of Acremonium strictum is maize (Pitt et al., 1993; Pitt and Hocking, 1997; Williams et al., 1992). It has recently been

isolated from black bean seeds in Argentina (Castillo et al., 2004) and raw cork (Quercus suber) in Spain (Alvarez-Rodriguez et al., 2002). Other records, including those of Acremonium spp. or the earlier name Cephalosporium acremonium, include wheat, barley and rice, bananas showing crown rot, fresh vegetables, peanuts, pecans, hazelnuts and walnuts, soybeans, frozen meat, salami and biltong (Pitt and Hocking, 1997). References. Gams (1971); Domsch et al. (1980).

60

5 Primary Keys and Miscellaneous Fungi

5.4 Genus Alternaria Nees: Fr. Alternaria produces large brown conidia with both longitudinal and transverse septa, borne from inconspicuous conidiophores, and with a distinct conical narrowing or ‘‘beak’’ at the apical end. These conidia are often formed in chains. Two other genera, Stemphylium Wallroth and Ulocladium Preuss, produce similarly septate conidia, but are uncommon in foods. Those of Ulocladium are roughly ellipsoidal, usually somewhat narrower at the base, and at most have small beaks. Both Alternaria and Ulocladium produce new conidia by lateral growth of the conidiophore from near the apex or through the beak area. Stemphylium conidia look like those of Ulocladium, but new conidia never form from a beak. New conidia are produced through the pore from which a conidium has already been produced. For a more detailed account of the distinctions among these three genera, see Simmons (1967).

Trichoconiella forms conidia similar to those of Alternaria, but with transverse septa only. This genus is associated with rice. Emory Simmons has spent a lifetime working with Alternaria and related genera and has recently revised the taxonomy of Alternaria comprehensively (Simmons, 2007). The simple notions of earlier authors, i.e. that Alternaria alternata is everywhere and that specific plants were colonised by specific pathogens, have been painstakingly replaced by a much more robust taxonomy. For example, Alternaria from citrus fruits may be Alternaria citri, but may also be one of 20 other species. The monograph by Simmons (2007) includes 276 species and is an essential laboratory manual for anyone who wishes to identify Alternaria species from a range of sources. Alternaria is not an easy genus for the inexperienced, however. Treatment here is confined to three common species only, with cereals as their major source.

Key to included species 1. Conidia produced in unbranched chains on DCMA Conidia produced in branched clusters on DCMA

Alternaria tenuissima (see A. alternata) 2

2. Conidia in unbroken chains, i.e. narrow hyphal elements between spores lacking; species occurring on wheat and many other substrates Primary conidia producing secondary conidia, i.e. conidia often separated by short hyphal lengths; species usually occurring on wheat

Alternaria alternata (Fr.) Keissl.

Fig. 5.2

Alternaria tenuis Nees

Colonies on CYA and MEA 50–70 mm diam, or covering the whole Petri dish, plane, of deeply floccose off-white to grey brown mycelium; reverse brown to nearly black. Colonies on G25N 10–15 mm diam, low and dense, olive brown or grey; reverse brown to almost black. At 58C, at least microcolonies, often colonies up to 4 mm diam. Usually no growth at 378C, occasionally colonies up to 10 mm diam, similar in appearance to those at 258C, or white. Colonies on DCMA 60–70 mm diam or covering the whole Petri dish, plane, floccose but not sparse, mycelium grey to dark grey; reverse grey to dark grey or black, less commonly bluish black. Conidia blown out from the apices of undistinguished conidiophores as short, irregularly branched

A. alternata A. infectoria

chains of up to 10 units, and then septating both laterally and longitudinally, with up to six transverse and two to three longitudinal or oblique septa, usually of clavate or pyriform shape overall, tapering towards the apices, forming a short beak, in culture usually 20–40  8–12 mm, with walls smooth to conspicuously roughened. Distinctive features. A. alternata is the most common saprophytic Alternaria species, occurring on a wide variety of sources, in contrast to other species considered here. On DCMA, conidia are produced more or less uniformly above the agar surface, in quite lengthy chains. The larger conidia have both longitudinal and transverse septa. Taxonomy. The first edition of this book noted that with acceptance of the 1821 starting point date for Alternaria nomenclature, A. alternata slowly became accepted as the correct name for this

5.4 Genus Alternaria

61

a

b

c

Fig. 5.2 Alternaria alternata (a) colonies on PDA and DCMA, 7 d, 258C ; (b) conidia in situ, bar ¼ 50 mm; (c) conidia, bar ¼ 20 mm

species. With the reversion to be the 1753 starting point date, the correct name again became A. tenuis. However, publications by Fries, 1821–1832, were later given special status under the ICBN, so A. alternata is once more the correct name. Further changes appear unlikely. Physiology. Optimum growth of Alternaria alternata is near 258C with minima variously reported as – 5 to 6.58C and maxima near 368C (Hasija, 1970; Domsch et al., 1980). The minimum aw for growth at 258C is 0.86 (Hocking et al., 1994). Optimal growth occurs at pH 4–5.4, and the pH range for growth is 2.7–8.0 (Hasija, 1970). A. alternata is able to grow in oxygen concentrations as low as 0.25%

(v/v) in N2, with growth rates being proportional to oxygen concentration (Follstad, 1966; Wells and Uota, 1970). The presence of the volatile compounds 3-methyl-1-butanol, 1-octen-3-ol and 3octanone in stored wheat grains is usually correlated with infection by A. alternata, amongst other fungi (Sinha et al., 1988). Mycotoxins. Alternaria alternata produces several toxins, of which the most important is tenuazonic acid (TA) (Logrieco et al., 2003). Tenuzoic acid is toxic to a wide range of plants and animals, particularly mice, chick embryos and chickens (Logrieco et al., 2003). Sorghum grains containing high levels of TA were associated with the human

62

haematological disorder known as onyalay (Bottalico and Logrieco, 1998). Less toxic compounds include alternariol (AOH) and alternariol monomethyl ether (AME), altenuene (AE) and altertoxin I, II and III (Logrieco et al., 2003). AOH and AME are usually found in combination and have some teratogenic and fetotoxic effects in pregnant mice (Bottalico and Logrieco, 1998). The production of one or more of these toxins has been reported in tomatoes (Bottalico and Logrieco, 1998), citrus fruits (Logrieco et al., 1990a), capsicum and melons (Bottalico and Logrieco, 2001), cereal grains (Logrieco et al., 1990b; Webley et al., 1997), rapeseed and olives (Logrieco et al., 2003), and apples (Robiglio and Lopez, 1995; Tournas and Stack, 2001). Maximum production of alternariol, its monomethyl ether and altenuene occurred at 258C and 0.98 aw (Magan et al., 1984). However, the optimum for tenuazonic acid production was reported as 0.90 aw at 258C (Etcheverry et al., 1994). Altertoxins I, II and III are produced by many Alternaria species, however, A. alternata is the most important producer. Altertoxins have received much attention due to their mutagenic activity, particularly that of altertoxin III, whose mutagenicity is approximately ten times lower than that of aflatoxin B1 (Bottalico and Logrieco, 1998). Ecology. Under the names Alternaria alternata and A. tenuis, this species has been reported from a very wide range of foods. However, the very broad species concept used by many authors is not accepted by Simmons (2007). The following reports undoubtedly include a number of other species. It is a major pathogen of fresh tomatoes (Harwig et al., 1979; Zitter and Wien, 1984; Pose et al., 2004), though other Alternaria species are probably involved here too. Spoilage of eggplants and peppers (Snowdon, 1991), apples (Combrink et al., 1985) and bananas stored under modified atmospheres (Wade et al., 1993) have been reported. Damage may occur also to a wide variety of fresh vegetables including beans, cauliflowers, cucumbers, melons, peas and potatoes (Webb and Mundt, 1978; Snowdon, 1991). A. alternata has been recorded frequently from a wide range of cereals, particularly wheat (Pitt et al., 1998b; Clear et al., 2005, Fakhrunnisa et al., 2006; Lugauskas et al., 2006 and references quoted in Pitt and Hocking, 1997), barley (Andersen et al., 1996; Fakhrunnisa et al., 2006; Medina et al., 2006 and references

5 Primary Keys and Miscellaneous Fungi

quoted in Pitt and Hocking, 1997), but also in rice, rapeseed, sunflower seed (Pozzi et al., 2005) and sorghum (Fakhrunnisa et al., 2006). In Australia and Southeast Asia, A. alternata has caused severe damage, spoilage and mycotoxin production in wheat and sorghum (Pitt et al., 1994; Webley et al., 1997 and our unpublished observations). This species has been recorded quite frequently from nuts, including peanuts, hazelnuts and pecans (see Pitt and Hocking, 1997). Other sources include soybeans, cold stored and frozen meat, biltong, bottled water and spices (Pitt and Hocking, 1997; Cabral and Fernadez Pinto, 2002). This species has occasionally been associated with gushing in beer (Niessen et al., 1992). Closely related species. Alternaria tenuissima (Kunze) Wiltshire differs from A. alternata by the formation of longer conidia (up to 60 mm) in unbranched chains (Simmons, 2007). The metabolite profiles of A. alternata and A. tenuissima are very similar. In a recent study, it was found that most A. tenuissima isolates examined produced alternariol, alternariol monomethyl ether and tenuzoic acid, whereas three of the A. alternata isolates produced altenuene, alternariol, alternariol monomethyl ether and altertoxin I (Andersen et al, 2002). Andersen and Frisvad (2004) reported A. tenuissima from mouldy tomatoes. In our experience this species is of common occurrence in cereals. References. Simmons (1967, 2007); Ellis (1971); Domsch et al. (1980).

Alternaria infectoria E.G. Simmons

Fig. 5.3

Colonies on CYA and MEA 60 mm diam or more, on CYA plane, low to moderately deep, mycelium dark grey to almost black, sometimes with a paler grey floccose overlay; reverse bluish black; on MEA moderately deep to deep, dense to floccose, of dark grey mycelium sometimes overlaid with lighter grey hyphae; reverse bluish black. On G25N, colonies 8–15 mm diam, of grey or greenish grey mycelium; reverse pinkish, grey or black. At 58C, at least germination, usually colonies of 2–5 mm diam formed. At 378C, colonies 5–25 mm diam, low to deep, grey, reverse dark. Colonies on DCMA 60 mm or more diam, usually with wide margins, low to subsurface,

5.4 Genus Alternaria

63

a

b

c

Fig. 5.3 Alternaria infectoria (a) colonies on PDA and DCMA, 7 d, 258C ; (b) conidia, bar ¼ 50 mm; (c) conidia, bar ¼ 20 mm

centrally usually sparse and floccose, aerial mycelium white to pale olive brown; reverse pale brown, brown or olive brown, sometimes darker in annular or irregular patches. Conidia on DCMA observed under the stereomicroscope characteristically formed in short, irregularly branched chains, forming clusters, with primary conidia borne from knobby hyphae, usually bearing secondary conidia borne from the tips, at the end of short (30–100 mm) hyphal lengths; conidia characteristically clavate (clubshaped) 20–50  7–14 mm, with up to 7 transverse septa and 1 or more longitudinal septa as well, or sometimes

irregularly septate, walls and septa thick and dark brown, and with walls sometimes conspicuously roughened. Distinctive features. After growth on DCMA, Alternaria infectoria is readily distinguished from A. alternata and other Alternaria species under the stereomicroscope. A. infectoria forms clusters of conidia borne in irregularly branching chains that, unlike those of A. alternata, have short hyphal segments between the conidia. Colonies on DCMA have low, wide margins, are less floccose than those of A. alternata and (apart from conidia) have paler colours.

64

Taxonomy. This species was commonly known as the Alternaria state of Pleospora infectoria until Simmons (1986) provided a binomial name, A. infectoria. He showed that its teleomorph does not belong in Pleospora, as had been widely accepted, and erected a new genus, Lewia Barr and Simmons, for it (Simmons, 1986). Simmons (1986) reported that Alternaria species related to A. infectoria have very rarely been seen to produce a teleomorph; however, on several occasions we have seen Lewia teleomorphs in isolates of A. infectoria grown on autoclaved wheat grains. Physiology. No physiological studies on this species have been reported. Mycotoxins. Alternaria infectoria is chemically very different from other Alternaria species (Andersen et al., 2002). A. infectoria does not produce any of the known Alternaria metabolites (Andersen and Thrane, 1996) and shares only a few metabolites with other Alternaria species (Andersen et al., 2002). We have confirmed the absence of any toxin production by this species (unpublished). Ecology. Alternaria infectoria is most commonly associated with grains, having been reported from small grains in the United States (Bruce et al., 1984), Australian wheat (Pitt et al., 1998b), Danish barley (Andersen et al., 1996) and Norwegian grains (Kosiak et al., 2004). The lack of other reports of

5 Primary Keys and Miscellaneous Fungi

A. infectoria more likely reflects failure by most investigators to differentiate Alternaria species, rather than true rarity. It seems probable that this species may be confined to small grains. References. Simmons (1986, 2007).

5.5 Genus Arthrinium Kunze Arthrinium, in earlier literature more commonly known by its synonym Papularia, is not a common genus in foods, but has occasionally caused spoilage. Arthrinium produces relatively large, darkwalled (but not black) conidia borne solitarily, both terminally and laterally, on short, narrow conidiophores. Two species are treated here, A. phaeospermum and the Arthrinium state of Apiospora montagnei, distinguished from each other by conidial size.

Arthrinium phaeospermum (Corda) M.B. Ellis

Fig. 5.4

Papularia sphaerosperma (Pers.) Hohn. ¨

Colonies on CYA and MEA covering the whole Petri dish, mycelium low or floccose, coloured white or grey, sometimes with conspicuous areas of pink,

Fig. 5.4 Arthrinium phaeospermum (a) colony on CYA, 7 d, 258C; (b) conidia, bar ¼ 10 mm

5.6 Genus Aureobasidium

darkening in age; reverse yellow or brown. Colonies on G25N 10–18 mm diam, of white mycelium. Sometimes germination at 58C. No growth at 378C. Reproduction by solitary conidia, blown out from the ends of, or from denticles on the sides of, short, narrow, sometimes sinuous conidiophores, themselves borne in clusters from mother cells on natural substrates, but often singly in culture; conidia circular in plan view but elliptical from the side, 8–12  5–7 mm, dark brown, smooth walled, often with a narrow, hyaline band around the longest periphery. Distinctive features. The conidium of Arthrinium is distinctive: solitary, dark-walled, circular in plan but elliptical from the side, and often with a hyaline peripheral band. Cultivation on DCPA and under lights may assist sporulation. In the present context, Arthrinium is distinguished from Nigrospora by the latter’s production of jet black conidia entirely devoid of ornamentation. Physiology. Conidia of Arthrinium appear to be highly heat resistant. In apple juice, conidia survived a pasteurising process of 888C for 1.5 min, and in water, heating at 1058C for 2.5 min (Anon, 1967). No experimental details, i.e., numbers heated or come-up time, were reported, however, and a decimal reduction time cannot be calculated. Even so, survival of a heat treatment at 888C indicates that A. phaeospermum is uncommonly heat resistant for a conidial fungus. Optimum germination of A. phaeospermum occurs when the conidia are suspended in sterile saline solution at 208C for 15 min before plating onto 2% malt extract agar (Agut and Calvo, 2004). The authors also noted that the viability of Arthrinium conidia was low, making this genus relatively rare in the environment. Mycotoxins. Arthrinium species are not known to produce mycotoxins, however, reports of secondary metabolites with antimicrobial properties have been found for some Arthrinium species (quoted in Agut and Calvo, 2004). Ecology. Arthrinium phaeospermum (reported as Papularia sphaerosperma) has been recorded as a cause of spoilage of pasteurised apple juice (Anon., 1967). It has also been reported on barley, wheat flour, rice, pecans and airline meals (see Pitt and Hocking, 1997). It was isolated from Morelo cherries (Olszak, 1994) and Iranian barley (Asgari et al., 2004). A. phaeospermum has also been recorded as a pathogen

65

on sweet potatoes (Ravichandran and Sullia, 1983). We isolated A. phaeospermum at low levels from paddy rice, mung beans, soybeans, black beans and cashews in Thailand (Pitt et al., 1993). Closely related species. The Arthrinium state of Apiospora montagnei Sacc., also known as Papularia arundinis (Corda) Fr., is similar to A. phaeospermum in all characters examined, except for the production of smaller conidia, 6–8 mm long. Data sheets at the International Mycological Institute, Egham, Surrey, UK, record its isolation from white flour and molasses; it has also been reported from wheat (Pelhate, 1968), barley (Flannigan, 1969) and cashews (Pitt et al., 1993). References. Ellis (1971); Domsch et al. (1980).

5.6 Genus Aureobasidium Viala and G. Boyer Growth of Aureobasidium isolates is at first yeast-like, but, while remaining very low and mucoid, colonies spread rapidly and turn black in patches. Microscopically, hyphae as well as budding yeast-like cells are present; the latter are actually conidia. The conidia are borne from small denticles (minute projections) directly from the hyphal walls or from short lateral protrusions on the hyphae; characteristically 2–4 denticles on one cell will produce conidia synchronously. There are more than 10 species (Hermanides-Nijhof, 1977), but only one, Aureobasidium pullulans, is of importance in foods (Fig. 5.5).

Aureobasidium pullulans (de Bary) G. Arnaud

Fig. 5.5

Dematium pullulans de Bary Pullularia pullulans (de Bary) Berkhout

Colonies on CYA and MEA 25–35 mm diam, low and mucoid, faintly pink, becoming grey to black in areas at 7–10 days; reverse in similar colours. Colonies on G25N 10–12 mm diam, similar to those on CYA. At 58C, microcolonies to colonies up to 3 mm diam. No growth at 378C. Conidia borne on denticles directly from hyphae or sometimes small lateral protrusions; conidia yeast-like, primary ones borne from the denticles, usually measuring 10–16  3–6 mm, and secondary

66

5 Primary Keys and Miscellaneous Fungi

a

b

c

Fig. 5.5 Aureobasidium pullulans (a) colonies on CYA and MEA, 7 d, 258C; (b, c) conidia, bar ¼ 10 mm

ones by budding from the primaries, 7–10  3–5 mm, not adhering to each other, smooth walled. Distinctive features. Within the present context, Aureobasidium pullulans is readily recognised by its distinctive low, mucoid, white then pink to black colonies and yeast-like conidia. Several other genera have a similar appearance (Hermanides-Nijhof, 1977; Beh, 2007) but they rarely occur in foods. Taxonomy. Earlier literature discusses this species under the names Dematium pullulans and Pullularia pullulans. Mycotoxins. Mycotoxins are not known to be produced. Physiology. The temperature range for growth of Aureobasidium pullulans has been reported as 2–358C, with an optimum of 258C (Skou, 1969), but some earlier data, together with its abundance

in low temperature habitats, suggests that some strains grow down to –58C (Michener and Elliott, 1964). It is of common occurrence in salterns (Butinar et al., 2005a), so is likely to be a xerophile. Heat resistance is very low (Skou, 1969). The ability of A. pullulans to grow on sugars used in yeast taxonomy has been reported by de Hoog and Yurlova (1994) and Beh (2007). Ecology. A ubiquitous saprophyte from all sorts of moist and decaying environments, Aureobasidium pullulans has been reported from a very wide range of foods, but only rarely as a cause of spoilage. Its prevalence in frozen foods is noteworthy, being the predominant mould isolated from blueberry, apple and cherry pies by Kuehn and Gunderson (1963). It was the most frequently isolated mould in Canadian icewine must, but played no role in fermentating

5.7 Genus Bipolaris

must (Subden et al., 2003). The surface of healthy grapes is predominately colonised by A. pullulans (Fleet, 2003a; Prakichaiwattana et al., 2004; Beh, 2007). A. pullulans has been reported in yoghurt in a survey of a Slovakian dairy plant (Pieckova et al., 2002) and has been associated with the spoilage of cold stored meat and cheese (see Pitt and Hocking, 1997). A. pullulans has been recorded in a variety of fresh foods and commodities including fresh vegetables, cabbage, strawberries, grapes, citrus and pasteurised orange juice (see Pitt and Hocking, 1997). Other records include shrimp, green olives, barley, wheat and flour, oats and nuts (see Pitt and Hocking, 1997). References. Hermanides-Nijhof (1977); Domsch et al. (1980); de Hoog and Yurlova (1994).

5.7 Genus Bipolaris Shoemaker In separating unrelated species from Helminthosporium, Shoemaker (1959) revised the genus Drechslera S. Ito and erected Bipolaris. The fact that species of Bipolaris (in the sense of Shoemaker, 1959) were associated with two distinct teleomorph types, and that correlated with morphological differences, led to segregation of Exserohilum from Bipolaris by Leonard and Suggs (1974). These genera produce long, large conidia with transverse septa only; the septa are thick and quite different from those in e.g. Trichoconiella. In Drechslera, conidia are cylindrical and germinate at any cell, while conidia of Bipolaris and Exserohilum are fusoid, i.e. gradually narrowing, and conidia germinate only from the end cells (bipolar germination). In Bipolaris, germ tubes develop roughly along the line of the conidium axis, while in Exserohilum, the germ tube is often offset from the axis and develops at an angle (Alcorn, 1983). Exserohilum species are apparently rare in foods, although E. rostratum (Drechsler) K.J. Leonard and Suggs has been reported from sorghum (Usha et al., 1994). The literature on these genera is voluminous and complex. The most useful publications are by Alcorn (1983) and Sivanesan (1987). One common species of Bipolaris, B. maydis, is treated here (Fig. 5.6).

67

Bipolaris maydis (Nishik. and C. Miyake) Shoemaker Fig. 5.6 Colonies on CYA and MEA 45–55 mm diam, plane, deeply floccose, of light to dark grey mycelium; reverse dark brown to dark blue black. On G25N, colonies 2–6 mm diam. At 58C, germination. At 378C, colonies 5–15 mm diam, usually low, dense and wrinkled, of grey mycelium, reverse deep blue black. Colonies on DCMA 30–40 mm diam, plane, of sparse to dense, floccose grey mycelium, reverse brown, grey or black. Conidia borne singly from nodes on knobby or geniculate mid to dark brown hyphae, usually slightly curved, (40–)50–80  12–16 mm, with six to nine inconspicuous septa and with smooth, dark, thick walls. Distinctive features. Large conidia, often slightly curved, of fusoid shape and with several thick transverse septa are characteristic of this genus. Conidial curvature and shape distinguish it from Drechslera, while conidial length and the presence of more than three, thick septa allow distinction from Curvularia. The species treated here is common on maize. Taxonomy. Some species of Bipolaris produce teleomorphs, classified in the genus Cochliobolus. Physiology. Species of Bipolaris sporulate significantly better when grown on substrates containing cellulose (Pratt, 2006). Mycotoxins. No significant mycotoxins are known. Ecology. We have isolated Bipolaris maydis from 1% of Thai maize kernels, and less frequently from Thai sorghum and mung bean samples (Pitt et al., 1993, 1994). It was also found at similar levels in commodities from Indonesia and the Philippines (Pitt et al., 1998a). Other records of Bipolaris in foods are rare. However, references to Drechslera species in foods often have referred to taxa correctly classified in Bipolaris, e.g. in rice, B. oryzae (Breda de Haan) Shoemaker; in barley, B. australiensis (M.B. Ellis) Tsuda and Uemaya, B. maydis and B. setariae (Sawada) Shoemaker; in peanuts, B. spicifera (Bainier) Subram.; in coriander seed, B. bicolor (Mitra) Shoemaker; and in avocados, B. setariae (see Pitt and Hocking, 1997). References. Alcorn (1983); Sivanesan (1987).

68

5 Primary Keys and Miscellaneous Fungi

a

b

c

Fig. 5.6 Bipolaris maydis (a) colonies on MEA and DCMA, 7 d, 258C; (b, c) conidiophores and conidia, bars ¼ 10 mm

5.8 Genus Botrytis P. Micheli: Fr. Botrytis is a common genus in the temperate zones, where it occurs mainly as a pathogen on a variety of plant crops. Vegetables and small berry fruits are particularly susceptible. Invasion may occur before maturity or postharvest, both in transport and in storage. Onions and other Allium species and grapes are the most susceptible crops. In the latter, it is notable that the disease is sometimes encouraged. Grapes affected by Botrytis, in this circumstance called ‘‘the noble rot’’, are used in the production of certain high quality sweet wines in France, Germany, Australia and other countries. Botrytis species are characterised by the production of conidia on pegs from spherical swellings. The most commonly encountered species in foods

is known as Botrytis cinerea, although it is probable that this name includes a group of related species rather than a single well defined taxon (Coley-Smith et al., 1980).

Botrytis cinerea Pers.

Fig. 5.7

Colonies on CYA and MEA covering the whole Petri dish, floccose, growth sometimes patchy or irregular, mycelium white, becoming grey to dark grey as conidiogenesis proceeds; reverse pale to grey. On G25N, colonies 10–18 mm diam, irregular in outline, floccose centrally or in patches, becoming grey; reverse grey. At 58C, colonies up to 5 mm diam produced, low and sparse. No growth at 378C.

5.8 Genus Botrytis

69

Fig. 5.7 Botrytis cinerea (a) colony on CYA, 7 d, 258C; (b) conidiophores and conidia, bar = 25 mm

Conidiophores borne from aerial hyphae, stipes of indeterminate length, each bearing terminally an irregular cluster of short branches, 10–30 mm long, with swollen spherical apices, 8–10 mm diam; conidia borne singly from these apices on denticles (small pegs), ellipsoidal, 8–12 mm long, smooth walled, not released at maturity. Distinctive features. Solitary conidia borne on denticles from terminally swollen, short branches are characteristic of Botrytis. Physiology. Growth has been reported at 0.93 aw (Snow, 1949) and at 0.90 aw (Jarvis, 1977). Lahlali et al. (2007) observed no growth of Botrytis cinerea at 0.93 aw in the presence of NaCl or at 0.89 aw in the presence of nonionic solute. Reported growth temperatures are rather variable, with minima from –2 to 58C or even 128C, maxima 28–358C, and optima of 22–258C (Domsch et al., 1980). B. cinerea will grow from pH 2–8 (Jarvis, 1977) and in O2 concentrations down to 1% (Follstad, 1966). The effect of low temperatures and controlled atmosphere storage on the growth of B. cinerea on susceptible fruits and vegetables has been reported (Reyes, 1990). B. cinerea produces pectinases (polygalacturonase) and the cell wall degrading enzymes cellulase and xylanase (El-Habbaa, 2003). Mycotoxins. Mycotoxin production has not been reported – fortunately, in view of this species role in speciality wine production.

Ecology. Botrytis cinerea is a virulent pathogen, the cause of rots in many kinds of fresh fruits (Snowdon, 1990). It is the most important cause of disease in grapes, both before harvest and in storage and the most common cause of spoilage in berries such as strawberries, blueberries, raspberries and blackberries (Snowdon, 1990; Tournas and Katsoudas, 2005), and also causes large losses in apples and pears, tomatoes, stone fruits and kiwi fruit (see Pitt and Hocking, 1997). A gel-like turbidity can be caused in raspberry juice, making filtration difficult (Will et al., 1992). For a review of many aspects of occurrence, pathogenicity and control, see Elad et al. (2004). Although vegetable spoilage is perhaps less important, the range of crops affected is very large, and includes asparagus, beans, cabbages, carrots, celery, melons and potatoes (Snowdon, 1991; Lugauskas et al., 2005; Tournas, 2005). It has also been isolated from a wide variety of dried or processed foods, but here its role is uncertain and it is probably present only as a contaminant. Other important species. Several other species of Botrytis are commonly occurring pathogens. Of interest here are B. allii Munn, B. byssoides Walker and the Botrytis state of Sclerotinia porri van Beyma, all of which cause spoilage of onions and leeks (Ellis, 1971; Snowdon, 1990). Botrytis allii grows optimally at 20–258C, with a minimum below 58C and a maximum near 358C. The optimum aw for growth is in excess of 0.99. Growth

70

5 Primary Keys and Miscellaneous Fungi

occurs down to 0.96 aw in media containing NaCl and 0.93 aw in media containing KCl or sucrose, or on onion leaves (Alderman and Lacy, 1984). References. Ellis (1971); Coley-Smith et al. (1980); Domsch et al. (1980); Elad et al. (2004).

5.9 Genus Chaetomium Kunze Chaetomium is the only genus of Ascomycetes which produces black ascocarps and which is commonly encountered in foods. The ascocarps in Chaetomium are perithecia, similar in appearance to cleistothecia, but having an opening (ostiole) through which the ascospores are released. There is a superficial resemblance to the genus Phoma, which produces black

conidiomata (pycnidia); ascospores in Chaetomium are often freed from the ascus before being discharged singly from the perithecium, and hence could be mistaken for conidia. Chaetomium is readily distinguished from Phoma by the presence of stout black or dark coloured hyphae attached to the perithecial walls, which are visible in culture under the low power microscope. Chaetomium species are notable as producers of cellulases and so commonly occur on wood and paper products. They are relatively common in some kinds of foods. C. globosum is the most frequently encountered species and is described here as representative of the genus. In addition descriptions of C. brasiliense and C. funicola, both quite common in tropical commodities, are provided here.

Key to Chaetomium species included here 1

Colonies on CYA at 378C much larger than at 258C Colonies on CYA at 378C smaller than at 258C or absent

2 (1)

Colonies on CYA and MEA at 258C less than 50 mm diam, ascospores 5–7 mm long Colonies on CYA and MEA at 258C usually covering the whole Petri dish, ascospores 8–10 mm long

Chaetomium brasiliense Bat. & Pontual

Fig. 5.8

Colonies on CYA and MEA 25–35 mm diam, on CYA low to moderately deep and dense, of white to grey mycelium; reverse sometimes pale, more commonly dark grey to almost black; on MEA low and sparse, dark grey, with some paler grey aerial hyphae; reverse dark grey. No growth on G25N or germination at 58C. Colonies at 378C covering the whole Petri dish, 65 mm or more diam, general appearance similar to those on CYA at 258C; reverse very dark grey to greyish black (Fig. 5.8). Reproductive structures perithecia not observed on CYA or MEA at 258C, but on CYA at 378C abundant, forming a deep, contiguous layer on the agar surface, most readily observed at colony margins, especially between colonies, surrounded by radiating long, stout hyphae, straight at first but at maturity curling into long corkscrew shapes, 75–100 mm diam, maturing in 7 days at colony centres; asci not seen, ascospores

C. brasiliense 2 C. funicola C. globosum

subspheroidal to broadly ellipsoidal, 6–8 mm long, brown, with smooth walls. Conidia not produced. Distinctive features. Unlike the other Chaetomium species considered here, C. brasiliense grows much more rapidly at 378C than 258C, and perithecia are formed in much greater numbers at the higher temperature. Perithecia are black, but under the stereomicroscope appear surrounded by grey hyphae. Ascospores are smaller than those of C. globosum. Physiology. No studies on physiology are known to us. However, the description above indicates that this species has a high temperature optimum for growth. Mycotoxins. Significant mycotoxins are not known to be produced. Ecology. We have isolated this species not infrequently from tropical sources. From Thailand, 1% of mung beans, black beans, paddy rice grains and cashew kernels, and 7–10% of soy bean and sorghum samples, but <1% of individual seeds, contained C. brasiliense (Pitt et al., 1993, 1994). Similar figures were obtained from Indonesian commodities (Pitt et al., 1998a). In samples from the Philippines, this species was found in maize, peanuts, soybeans, mung beans,

5.9 Genus Chaetomium

71

Fig. 5.8 Chaetomium brasiliense (a) colony on CYA, 7 d, 258C; (b) perithecium in situ, bar ¼ 25 mm; (c) asci, bar ¼ 10 mm; (d) ascospores, bar ¼ 5 mm

black pepper, paddy and milled rice, all near 1% of total particles examined (our unpublished data). Reference. Von Arx et al. (1986).

Chaetomium funicola Cooke

Fig. 5.9

Colonies on CYA 20–25 mm diam, plane, of low and sparse white mycelium, sometimes pale grey centrally, enveloping abundant developing perithecia, reverse pale or greyish brown. Colonies on MEA 35–40 mm diam, plane, sparse, low to somewhat floccose, mycelium white or grey, enveloping developing perithecia; reverse pale to greyish olive or pale brown. On G25N, usually no growth. At 58C, no germination. At 378C, usually no growth.

Perithecia borne in a layer on the agar surface, 120–160 mm diam, surrounded by almost straight, dark walled, rough to spiky hyphae; asci maturing in 14 days, breaking down immediately, ascospores ellipsoidal, 5–7  3–5 mm, grey or brown, sometimes with one or two internal oil droplets visible, often collapsing in age, smooth walled. Distinctive features. Chaetomium funicola grows only slowly at 258C and usually not at 378C. Ascospores are smaller than those of C. globosum. Physiology. No studies are known to us. Domsch et al. (1980) indicated that perithecial production is maximal at about 208C, indicating that this species is better adapted to low than high temperatures.

72

5 Primary Keys and Miscellaneous Fungi

Fig. 5.9 Chaetomium funicola (a) colonies on CYA and MEA, 7 d, 258C; (b) perithecium in situ, bar ¼ 50 mm; (c) ascospores, bar ¼ 5 mm

Mycotoxins. Mycotoxin production has not been reported. Ecology. Despite its low temperature optimum, we repeatedly isolated Chaetomium funicola from tropical commodities. It was present in 23% of Indonesian soybean samples, ranging up to 50% infection in some samples, and in 3% of beans overall (our unpublished data). Lower levels of infection (1% or more overall) were observed in peanuts, cashews, copra, mung beans and sorghum from Thailand (Pitt et al., 1993, 1994), in mung beans and maize from Indonesia, and paddy rice from the Philippines (Pitt et al., 1998a and our unpublished data). References. Domsch et al. (1980); von Arx et al. (1986).

Chaetomium globosum Kunze

Fig. 5.10

Colonies on CYA and MEA covering the whole Petri dish, on CYA low and sparse, of scanty white mycelium and conspicuous though usually sparse black perithecia, ca. 0.2 mm diam; on MEA growth more dense but still low, coloured grey or greenish black from hyphae enveloping abundant perithecia; reverse on both media usually brown. On G25N, colonies less than 5 mm diam produced. No growth at 58C. At 378C, colonies usually 20–30 mm diam, of white mycelium. Reproductive structures perithecia, black, 150–200 mm diam, with numerous stout, dark hyphae appended; ascospores produced after 1–2 weeks,

5.10 Genus Chrysonilia

73

Fig. 5.10 Chaetomium globosum (a) colony on CYA, 7 d, 258C; (b) perithecium in situ, bar ¼ 100 mm; (c) ascospores, bar ¼ 5 mm

spheroidal, broadly ellipsoidal or apiculate, commonly 8–10 mm long, smooth walled. Conidia not generally produced. Distinctive features. In contrast to Chaetomium brasiliense, C. globosum grows much more rapidly at 258C than 378C. Ascospores are larger than those of C. funicola. Physiology. Chapman and Fergus (1975) reported that Chaetomium globosum ascospores germinated from a minimum temperature of 4–108C to a maximum of 388C, and most rapidly at 24–388C. They also reported germination over the whole pH range tested, 3.5–7. Heat resistance was low: 1% of ascospores survived 10 min at 558C, but none survived at 578C for a similar period. Ahammed et al. (2005) observed maximum sporulation at 288C and pH 5.0 on a medium containing sucrose and thiamine. C. globosum can grow down to 0.94 aw in soil (Kouyeas, 1964). Mycotoxins. Chaetomium globosum produces chaetoglobsins A and C, moderately toxic compounds reported to be produced in building materials (Fogle et al., 2007). Their relevance to foods is unknown. Ecology. This species has been isolated from a variety of commodities, particularly wheat, barley, rice, beans and soybeans. Chaetomium globosum has also been recorded in nuts (see Pitt and Hocking, 1997) and spices (Mandeel, 2005). We isolated it

frequently from tropical commodities: from 12 to 17% of maize samples, 14 to 32% of mung bean samples and 33 to 35% of soybean samples from Thailand, Indonesia and the Philippines, with overall infection rates of 1–4% of all particles examined. Relatively high infection levels were also encountered in Thai cashews, copra and paddy rice. Kemiri (candle) nuts from Indonesia were highly infected: 37% of samples, up to 40% in infected samples and 9% of nuts overall (Pitt et al., 1998a). This species has not been reported to cause food spoilage, although it has recently been implicated in causing disease in pears in Egypt (Ismail and Abdalla, 2005). References. Domsch et al. (1980); von Arx et al. (1986).

5.10 Genus Chrysonilia Arx Chrysonilia was erected by von Arx (1981a) to accommodate Monilia sitophila, for many years known as ‘‘the red bread mould’’. Chrysonilia sitophila is characterised by very rapid growth and the production of conidia in chains, cut off from the apices of undifferentiated hyphae. This species is readily recognisable in Petri dish culture: sparse, white to pink mycelium spreads rapidly across the dish and up the walls, then within 3 days sheds

74

5 Primary Keys and Miscellaneous Fungi

Fig. 5.11 Chrysonilia sitophila (a) colony on CYA, 7 d, 258C; (b) conidia, bar ¼ 10 mm

enormous numbers of orange conidia outside the dish. This attribute makes it a particularly troublesome laboratory contaminant. Cultivation in Petri dishes is therefore not recommended: observation of these characters in a culture tube is sufficient for positive identification. The teleomorph of this species is Neurospora sitophila, a well-known Ascomycete which has been of great value in genetic studies. It is not found in foods, however.

Chrysonilia sitophila (Mont.) Arx

Fig. 5.11

Monilia sitophila (Mont.) Sacc.

Teleomorph: Neurospora sitophila Shear and B.O. Dodge Colonies in 28 ml McCartney bottles or test tubes distinguished by pale pink floccose growth, filling the entire bottle, then turning salmon, first at the bottle neck or tube plug and subsequently throughout, as sporulation occurs. On CYA and MEA, colonies covering the whole Petri dish, mycelium pink, reaching the lid in tufts or patches and all around the rim, producing vast numbers of salmon conidia at or near the rim and sometimes for several millimetres beyond it, and shedding them profusely; reverse salmon or pink. On G25N, colonies up to 30 mm diam produced,

low, dense and mucoid. No growth at 58C. At 378C, colonies covering the whole Petri dish, similar to at 258C. Reproductive structures arthroconidia, cut off in succession from the apices of branching hyphae; conidia at maturity variable in size and shape, spherical to ellipsoidal, pale orange, 6–15 mm diam, with thin smooth walls. Distinctive features. See remarks above. Physiology. According to Panasenko (1967), Chrysonilia sitophila is capable of growth down to 0.88–0.90 aw. Mycotoxins. No mycotoxins have been recorded for this species. Ecology. For long a common sight in bakeries and on bread, Chrysonilia sitophila is less commonly encountered now, but can still be a great source of trouble as a persistent contaminant in laboratories (see Chapter 4). It has sometimes been reported from other foods: pastries, hazelnuts, beans and meat products (see Pitt and Hocking, 1997). We isolated it frequently from Thai cassava, where it infected 36% of all pieces examined, and Philippine peanuts, where it infected up to 90% of nuts in some samples, and was present in 1% of nuts overall. It was also present in 1% of all Indonesian peanuts, sorghum and maize grains examined (Pitt et al., 1998a). Infection rates in maize and peanuts ranged up to 40% in some samples (our unpublished data). We also found it at lower levels in Thai soybeans, black beans and

5.11 Genus Cladosporium

75

cashews (Pitt et al., 1994). Recently, C. sitophila was reported in the mycoflora of cork slabs (Oliveira et al., 2003; Pereira et al., 2006). Reference. Von Arx (1981a).

5.11 Genus Cladosporium Link Cladosporium is a very commonly isolated genus and is often dominant in studies of airborne microflora. Species occur both as saprophytes and as plant pathogens. Conidia of Cladosporium species are particularly well adapted to aerial dispersal, being small, dry, heavily pigmented and apparently highly resistant to sunlight. The genus is highly diverse, and includes many species not yet characterised (Crous et al., 2007). In culture, Cladosporium is readily recognised. Colonies are 15–40 mm in diameter, low, dense and velvety, and coloured olive. Colony reverses on CYA are often a deep iridescent blue black, uncommon among slowly growing genera. Reproductive structures are fragile, tree-like conidiophores, which branch irregularly by budding from the youngest cells. The structures can be seen by examination of colonies under the

stereomicroscope. They disintegrate partially or totally in wet mounts, leaving masses of conidia, which may show buds or bud scars. In shape, but not colour, the conidia often resemble yeast cells; however, walls are thick, coloured olive and often roughened. Cladosporium species occur as pathogens on fresh fruit and vegetables (Snowdon, 1990, 1991) and may cause spoilage of strawberries (Beneke et al., 1954; Dennis et al., 1979) or tomatoes (Harwig et al., 1979). On other foods, Cladosporium species usually occur as contaminants rather than as spoilage fungi. However, all common species grow at temperatures near 08C, and Cladosporia have been reported to cause spoilage of chilled meats, cheese and other refrigerated commodities from time to time (see Pitt and Hocking, 1997). A major taxonomic revision of Cladosporium, its teleomorphs and related genera has recently been published (Crous et al., 2007). In overall terms, the taxonomy of the common species treated here has been unaffected. The most frequently reported saprophytic species is Cladosporium herbarum, but C. cladosporioides and C. sphaerospermum are also ubiquitous. In addition, C. macrocarpum is described here.

Key to Cladosporium species included here 1

Single celled conidia small, less than 4 mm wide Single celled conidia often exceeding 4 mm wide

2 (1)

Single celled conidia ellipsoidal or apiculate (lemon-shaped) A high proportion (greater than 40%) of single-celled conidia roughly spherical or pyriform

3 (1)

Conidia 4–6 mm wide Conidia commonly exceeding 6 mm wide

Cladosporium cladosporioides (Fresen.) G.A. de Vries Fig. 5.12

Colonies on CYA and MEA 25–40 mm diam, low and dense, lightly wrinkled or plane, surface velutinous or lightly floccose; conidia abundant, coloured olive; reverse bluish grey. Colonies on G25N 5–12 mm diam, plane, sometimes centrally raised, velutinous, coloured as on CYA; reverse bluish black. At 58C, colonies usually 1–2 mm diam, occasionally only germination occurring. No growth at 378C.

2 3 C. cladosporioides C. sphaerospermum C. herbarum C. macrocarpum

Conidiophores in situ dendritic (tree-like), closely packed, with stipes bearing branching structures of acropetally produced cells, all functioning as conidia at maturity and separating in liquid mounts; conidia heavy walled, pale olive brown, larger ones non- or singly septate, 10–30  2–5 mm, smooth walled, smaller ones nonseptate, ellipsoidal to apiculate, 3–7  2–4 mm, with walls smooth to finely roughened. Distinctive features. Cladosporium cladosporioides produces smaller conidia than C. herbarum and C. macrocarpum; unlike C. sphaerospermum,

76

5 Primary Keys and Miscellaneous Fungi

Fig. 5.12 Cladosporium cladosporoides (a) colonies on CYA and MEA, 7 d, 258C; (b) fruiting structures in situ, bar = 50 mm; (c) conidiophores, bar = 5 mm; (d) conidia, bar = 5 mm

the majority of conidia are ellipsoidal. Growth on CYA and MEA is much faster than that of C. sphaerospermum. Physiology. Gill and Lowry (1982) reported a minimum growth temperature of –58C for Cladosporium cladosporioides; the maximum is near 328C (Domsch et al., 1980). It is capable of growth down to 0.86 aw at 258C (Hocking et al., 1994) and is relatively resistant to microwave heating (Dragoni et al., 1990). The minimum inhibitory concentration of sorbic acid is 160 mg/kg at pH 5 (Skirdal and Eklund, 1993). Mycotoxins. This species is not known to produce mycotoxins.

Ecology. Cladosporium cladosporioides has been isolated from a very wide variety of foods, including wheat and flour, barley, rice, sorghum, fresh and frozen meat and fresh vegetables (see Pitt and Hocking, 1997). It occurred on 16% of 74 dried fish samples from Indonesia (Wheeler et al., 1986), and was common on Egyptian walnuts and hazelnuts (Abdel-Hafez and Saber, 1993). It has also been found in peanuts (El-Magraby and ElMaraghy, 1988). Cladosporium cladosporioides is not as common a cause of spoilage in fresh fruit as C. herbarum, but can cause rots of raspberries and melons and sooty discolouration of bananas (Snowdon,

5.11 Genus Cladosporium

1990). Because of its psychrophilic nature, C. cladosporioides causes spoilage of refrigerated foods such as cheese (Northolt et al., 1980) and meat (Gill et al., 1981). It was the dominant fungus causing losses in vacuum packed Australian cheese blocks (Hocking and Faedo, 1992). Cladosporium cladosporioides is also very common in major Southeast Asian commodities: we isolated this species from 33% of mung bean, 14% of peanut, 13% of soybean and 10% of maize samples from the Philippines and 21% of soybeans and

77

14% of mung bean samples from Indonesia (Pitt et al., 1998a and our unpublished data). References. De Vries (1952); Ellis (1971); Domsch et al. (1980).

Cladosporium herbarum (Pers.) Link Fig. 5.13 Colonies on CYA and MEA 18–32 mm diam, velutinous to lightly floccose, plane or slightly wrinkled, coloured olive; reverse olive grey to dark greenish grey. Colonies on G25N 5–10 mm diam, low and

Fig. 5.13 Cladosporium herbarum (a) colonies on CYA and MEA, 7 d, 258C; (b) fruiting structures in situ, bar ¼ 50 mm; (c) conidiophores, bar ¼ 5 mm; (d) conidia, bar = 5 mm

78

sparse to deep and dense, coloured as on CYA or paler. At 58C, colonies usually 1–2 mm diam; occasionally only germination. No growth at 378C. Conidiophores in situ sparsely and irregularly branched dendritic structures, borne on long, dark stipes, separating in fluid mounts; conidia ellipsoidal to cylindroidal, extremities sometimes irregular due to bud scars, usually nonseptate but larger cells with one to two septa, commonly 8–15(–20)  4–6 mm, pale brown, with densely roughened walls. Distinctive features. The conidia of Cladosporium herbarum are 4–6 mm wide, with distinctly roughened walls; conidia of C. macrocarpum are 6–9 mm wide and have finely roughened walls. Taxonomy. It was concluded by Dugan and Roberts (1994) that Cladosporium herbarum represented part of a continuum with C. macrocarpum, while Ho et al. (1999) considered that C. macrocarpum was a variety of C. herbarum. However, in a major study, Schubert et al. (2007) have kept these two species. They indicated that C. herbarum as commonly used includes several other species, but the concept of C. herbarum used here is still valid. The teleomorph of C. herbarum is Davidiella tassiana (De. Not.) Crous and U. Braun (Schubert et al., 2007). This is not usually seen in pure culture. Physiology. Growth of this species has been reported down to 0.88 aw (Snow, 1949) and down to –108C (Joffe, 1962). The maximum growth temperature is 28–328C (Domsch et al., 1980). This species can grow and sporulate in an atmosphere containing 0.25% oxygen (Follstad, 1966). Mycotoxins. No mycotoxins are known to be produced. Ecology. Because Cladosporium herbarum has been the best known Cladosporium name for a long time, it seems likely that some reports of this species really refer to other Cladosporia, especially C. cladosporioides, in our experience the more common species in foods. For example, Waghray et al. (1988) reported C. herbarum as common on Indian rice, but according to Pitt et al. (1994, 1998a and unpublished), the common Cladosporium species on Southeast Asian rice is C. cladosporioides. In contrast to C. cladosporioides, C. herbarum causes spoilage of fresh fruits and vegetables: rots in fresh yams (Dioscorea sp.; Adeniji, 1970a), stored apples (Snowdon, 1990; Kalafatoglu and Karapinar, 1991), stone fruit, tomatoes, melons, grapes (Snowdon, 1990, 1991; Camili and Benato, 2005), passionfruit

5 Primary Keys and Miscellaneous Fungi

(Ribeiro and Dias, 2005) and pawpaw (Echerenwa and Umechuruba, 2004). Its common occurrence on fresh apples can lead to contamination of apple juice and high acid fruit based products. As a psychrophile, C. herbarum can cause ‘‘black spot’’ spoilage of meat in cool stores (Gill et al., 1981), of cheese during ripening (Gueguen, 1988) and in storage under vacuum (Hocking and Faedo, 1992). It has been isolated from fresh and frozen meat and processed meat products (see Pitt and Hocking, 1997). C. herbarum has also been isolated from eggs, peanuts, hazelnuts, walnuts, cereals, chickpeas, soybeans and frozen fruit pastries (see Pitt and Hocking, 1997; Kuehn and Gunderson, 1963; Arya, 2004). Growth on dough carrying machinery (Gradel and Mu¨ller, 1985) and on wine corks (Daly et al., 1984) has been reported. References. Dugan and Roberts (1994); Ho et al. (1999); Schubert et al. (2007).

Cladosporium macrocarpum Preuss

Fig. 5.14

Colonies on CYA 18–25 mm diam, velutinous to lightly floccose, plane or slightly wrinkled, coloured olive to dark olive; reverse bluish grey. Colonies on MEA 16–24 mm diam, low to floccose, pale olive to olive; reverse olive grey to dark olive grey. Colonies on G25N 3–8 mm diam, low and sparse to deep and dense, coloured as on MEA or paler. At 58C, colonies usually 1–2 mm diam; occasionally germination only or no growth observed. No growth at 378C. Conidiophores as seen under the stereomicroscope long and dark, with small clusters of conidia borne in short, irregularly branched chains; conidia borne from irregularly located pores on geniculate (‘‘knee-like’’) swellings, ellipsoidal or short cylindroidal, with rounded ends, (10–)15–20  6–9 mm, usually aseptate, with finely roughened brown walls. Distinctive features. Colonies are similar to those of Cladosporium herbarum, but of somewhat paler colour. Conidia are considerably larger than those of C. herbarum or of the other Cladosporium species described here. Taxonomy. From an examination of 32 isolates from cherries in Washington state, USA, Dugan and Roberts (1994) concluded that Cladosporium macrocarpum and C. herbarum represented part of a continuum and that C. macrocarpum was a

5.11 Genus Cladosporium

79

Fig. 5.14 Cladosporium macrocarpum (a) colonies on CYA and MEA, 7 d, 258C; (b) fruiting structures in situ, bar ¼ 50 mm; (c) conidiophores, bar ¼ 5 mm; (d) conidia, bar ¼ 5 mm

synonym. Ho et al. (1999) considered that C. macrocarpum was a variety of C. herbarum. However, in a major study, Schubert et al. (2007) kept the two species separate. Physiology. No reports on the physiology of this species have been located. Properties should be similar to those of Cladosporium herbarum, to which it is closely related. Mycotoxins. No mycotoxins have been reported. Ecology. Cladosporium macrocarpum is uncommon in foods. Abdel-Kader et al. (1979) and Petters et al. (1988) both reported it from barley, the

latter as one of the principal fungi found in a large survey. Dugan and Roberts (1994) reported frequent occurrence on Bing cherries in the United States (see Taxonomy). It occurs on hazelnuts and walnuts in Egypt (Abdel-Hafez and Saber, 1993). We have isolated this species at a low frequency from cashews in Thailand (Pitt et al., 1993) and mung beans in the Philippines (our unpublished data). It was also reported on fresh asparagus (Kadau et al., 2005). References. Dugan and Roberts (1994); Ho et al. (1999); Schubert et al. (2007).

80

Cladosporium sphaerospermum Penz. Fig. 5.15 Colonies on CYA 18–25 mm diam, low and dense, usually plane, surface velutinous; conidia abundant, coloured olive to dark olive; reverse bluish grey. Colonies on MEA 12–20 mm diam, of similar appearance to those on CYA, sometimes lighter in colour. Colonies on G25N 8–12 mm diam, plane or centrally raised, velutinous, coloured as on CYA; reverse almost black. At 58C sometimes germination. No growth at 378C. Conidiophores under the stereomicroscope dendritic (tree-like), closely packed, with stipes bearing branching structures of acropetally produced cells,

5 Primary Keys and Miscellaneous Fungi

all functioning as conidia at maturity, and separating in liquid mounts; conidia heavy walled, pale olive brown, larger ones 10–30  2–4 mm, only occasionally septate, smooth walled, smaller ones nonseptate, subspheroidal, ogival or apiculate, 4–6(–10)  3–4 mm, with walls smooth to definitely roughened. Distinctive features. Similar to Cladosporium cladosporioides in most respects, C. sphaerospermum is distinguished by slower growth on CYA and MEA and by production of a high proportion (40% plus) of more or less spherical conidia. Taxonomy. Several new species very similar to Cladosporium sphaerospermum were described by Zalar et al. (2007), but none was of common occur-

Fig. 5.15 Cladosporium sphaerospermum (a) colonies on CYA and MEA, 7 d, 258C; (b) fruiting structures in situ, bar ¼ 50 mm; (c,d) conidiophores and conidia (slide culture), bar ¼ 10 mm; (e) conidia, bar ¼ 5 mm

5.12 Genus Colletotrichum

rence. The concept of C. sphaerospermum as used here remains unchanged. Physiology. Cladosporium sphaerospermum has an optimum aw for growth near 0.97 at 258C. This species is a xerophile, able to germinate and grow slowly at 0.815 aw (Hocking et al., 1994). No other physiological studies have been reported. Mycotoxins. No mycotoxins are known to be produced. Ecology. Although less common than Cladosporium cladosporioides, C. sphaerospermum has been isolated from a wide range of foods, including barley and peanuts in Egypt, US pecan nuts, European meat products, spoiled cheese in Europe and Australia, stored apple fruit in Turkey (see Pitt and Hocking, 1997) and carbonated soft drinks in Nigeria (Odunfa, 1987) and Argentina (Ancasi et al., 2006) . We have isolated this species at low frequency from peanuts from Thailand (Pitt et al., 1993), rice and soybeans from the Philippines, and maize and mung beans from Indonesia (Pitt et al., 1998a). References. Zalar et al. (2007).

5.12 Genus Colletotrichum Corda Colletotrichum is a genus with many species, widespread as plant pathogens. In this genus, conidia are borne inside an acervulus, a structure with a more or less closed lid which eventually ruptures. Acervuli are readily seen on the agar surface with the unaided eye or by low power microscope. Conidia of Colletotrichum are single celled and hyaline or brightly coloured. They may be cylindrical or pointed, straight or curved. A number of Colletotrichum species cause spoilage of fresh fruits and vegetables. Examples include C. acutatum J.H. Simmonds on strawberries, C. circans (Berk.) Voglino on leeks and onions, C. coccodes (Wallr.) S. Hughes on potatoes, tomatoes and eggplants, C. higginsianum Sacc. on cabbage and other cucurbits, C. lindemuthianum (Sacc. and H. Magn.) Broome and Cavara on beans and C. musae (Berk. and Curtis) Arx on bananas (Snowdon, 1990, 1991). One species, Colletotrichum gloeosporioides, a common cause of spoilage of fruits, especially from the tropics, is treated here as an example of this rather ill-defined though important genus. In

81

the opinion of Sutton (1980), C. gloeosporioides as currently accepted is an agglomerate of several species with a similar appearance. The species description given below is taken from a limited number of isolates and undoubtedly does not cover the full range of variation.

Colletotrichum gloeosporioidides (Penz.) Sacc.

Fig. 5.16

Teleomorph: Glomerella cingulata (Stonemason) Spauld. & Schrenk. On CYA and MEA, colonies 60 mm diam or more, often covering the whole Petri dish, with a dense basal layer of hyphae and conidial fruiting bodies (acervuli) overlaid by areas of floccose white, orange or grey mycelium; acervuli up to 500 mm long, pale, grey or orange; reverse with pale grey or orange areas. On G25N, colonies 2–5 mm diam, pale or black. At 58C, no growth to germination. No growth at 378C. Reproductive structures flat, lidded acervuli, opening irregularly, containing a single closely packed layer of phialides, of irregular dimensions; conidia borne singly, cylindroidal, with rounded ends, nonseptate, 12–18  3–3.5 mm, hyaline and smooth walled. Distinctive features. In the present context, Colletotrichum gloeosporioides is distinguished by producing conidia in acervuli and by pathogenicity on tropical fruits; conidia are aseptate cylinders with rounded ends, 12–18 mm long. Taxonomy. This species as currently delimited is probably too broadly defined (Sutton, 1980). The teleomorph of Colletotrichum gloeosporioides, Glomerella cingulata (Stonemason) Spauld. and Schrenk., is not found in foods. Physiology. Colletotrichum gloeosporioides was insensitive to storage in 10–13% CO2 in the presence of 3–7% O2 (Wade et al., 1993). Mycotoxins. Mycotoxins are not known to be produced. Ecology. According to Sutton (1980), the International Mycological Institute Herbarium has records of Colletotrichum gloeosporioides from 470 different host genera – a remarkable host range, even taking into account that several biological species may be involved.

82

5 Primary Keys and Miscellaneous Fungi

Fig. 5.16 Colletotrichum gloeosporioides (a) colonies on CYA and MEA, 7 d, 258C; (b) immature conidiophores, bar = 10 mm; (c) conidia, bar = 5 mm

Colletotrichum gloeosporioides and closely related species cause anthracnoses of tropical fruits, including avocados, bananas, pawpaws (papayas) and mangoes, and of temperate fruits, including apples, stone fruits and grapes. Colletotrichum species cause damage to bananas (C. musae), strawberries (C. acutatum), grapefruit, tomatoes and other fruits (Snowdon, 1990; Pitt and Hocking, 1997). Anthracnoses on fruit are dark, relatively dry, shrunken skin blemishes which expand rapidly as the fruit ripens. Except in advanced stages, the blemishes are only skin deep, and the fruit is edible, if unsightly. Advanced lesions may develop pinkish masses of conidia. Control in bananas is generally possible with benzimidazole or similar fungicides; in mangoes, pawpaws and avocados, hot water dips, with or without fungicide, can be beneficial (Smoot

and Segall, 1963; Hall and Scott, 1977). Gamma radiation can be used to control C. gloeosporioides in pawpaws (papayas) (Cia et al., 2007). Colletotrichum species were isolated from 40% of paddy rice samples examined from Thailand (Pitt et al., 1994). C. dematium was found at low levels in several Southeast Asian commodities, including soybeans, cowpeas, peanuts and black pepper (Pitt et al., 1998a). References. Von Arx (1957); Mordue (1971); Sutton (1980); Gunnell and Gubler (1992).

5.13 Genus Curvularia Boedijn In Curvularia, conidia are long and ellipsoidal, with three to four transverse septa. As the name implies, conidia are often curved due to an asymmetrically

5.13 Genus Curvularia

83

swollen central cell. Most species are plant pathogens. The species most common in foods is

Curvularia lunata; Curvularia pallescens is described here as a second foodborne species.

Key to Curvularia species included here 1

Mycelium on CYA and MEA grey to dark grey; reverse often blue black; conidia with central cells dark-walled Mycelium on CYA and MEA pale to mid grey; reverse brown; all cells in conidia with uniform wall colour

Curvularia lunata (Wakker) Boedijn

Fig. 5.17

Teleomorph: Cochliobolus lunatus R.R. Nelson & F.A. Hassis On CYA and MEA, colonies at least 60 mm diam, often covering the whole Petri dish, usually deep, moderately dense and floccose, mycelium off-white to grey, often approaching black, in age sometimes developing orange or salmon coloured areas; reverse usually grey to bluish black, sometimes with areas of salmon. On G25N, colonies 5–15 mm diam, low and dense, grey to black with reverse similar. At 58C, usually germination. At 378C, colonies (5–)20–40 mm diam, of similar appearance to those at 258C. Colonies on DCMA 55–65 mm diam or covering the whole Petri dish, plane, sparse, velutinous, reverse brown or dark grey. Conidia, best seen in mounts from growth close to the agar surface on MEA or on DCMA, borne from pores along the sides of short knobby

C. lunata C. pallescens

conidiophores, elongate, smooth walled, with three septa, almost always curved at an asymmetric cell third from the base, 16–25(–30)  8–14 mm, end cells pale brown, central cells darker. Distinctive features. Curvularia lunata is the species of Curvularia commonly isolated from foods. It is distinguished from C. pallescens by darker colonies, a blue black reverse, and nonuniform pigmentation in conidial walls. Conidial production usually occurs on MEA, but culturing on DCMA and/or under lights may assist recognition if MEA plates are sterile. Taxonomy. When grown in pairs, some isolates of this species mate to produce an ascomycetous state, Cochliobolus lunatus R.R. Nelson and F.A. Haasis. This state is not encountered in agar culture of single isolates. Physiology. Curvularia lunata was able to germinate at 0.86 aw, but only grew down to 0.89 aw at 258C (Hocking et al., 1994). Mycotoxins. No reliable reports of mycotoxin production by this species are known to us.

Fig. 5.17 Curvularia lunata (a) colony on CYA, 7 d, 258C; (b, c) conidiophore and conidia, bar = 10 mm

84

Ecology. As Curvularia lunata is primarily an invader of monocotyledon plants (Domsch et al., 1980), the most common food sources are cereals: records include rice, barley, wheat, maize and sorghum (Pitt and Hocking, 1997; Mtisi and McLaren, 2003; Echemendia, 2005, Fakhrunnisa et al., 2006). It has been reported as one cause of spoilage of fresh tomatoes in Nigeria (Muhammad et al., 2004) and has also been found on litchi fruit (Wells et al., 1981), soybeans (Ahammed et al., 2006), hazelnuts and walnuts (Abdel-Hafez and Saber, 1993), peanuts and spices (see Pitt and Hocking, 1997). In Thailand, Curvularia lunata is a major invader of sorghum: we isolated it from 56% of samples and 8% of all grains examined. It was also prevalent in paddy rice samples, where 28% of samples and 3% of all grains examined were infected (Pitt et al., 1994). It was found in 9–10% of paddy rice samples from Indonesia and the Philippines (Pitt et al., 1998a and unpublished data). Additional variety. Curvularia lunata var. aeria (Bat.) M.B. Ellis is a floccose taxon accepted as a variety of C. lunata. Colonies of C. lunata var. aeria on CYA 50–65 mm diam, plane, floccose, of pale to mid grey mycelium, reverse deep blue black. Colonies on MEA covering the whole Petri dish, plane, deeply floccose, especially so at colony to colony and colony to Petri dish junctions, mycelium white to orange grey, reverse yellow brown, dark brown or blue black. Colonies on G25N 4–6 mm diam, of dark grey mycelium. No growth at 58C. At 378C, colonies 50–65 mm

5 Primary Keys and Miscellaneous Fungi

diam, similar to those on CYA at 258C, sometimes less floccose, reverse dark blue black. Colonies on DCMA 55–65 mm diam, plane, sparse, floccose, especially at colony junctions, reverse brown or dark grey. Conidia are similar to those of C. lunata. The variety is distinguished from Curvularia lunata var. lunata by a more floccose growth habit and faster growth at 378C. It occurs in similar foods and commodities to C. lunata, the highest level we have encountered being 2% of all Thai rice grains and 1% of all Thai sorghum grains examined (Pitt et al., 1994). References. Ellis (1971); Domsch et al. (1980); Sivanesan (1987).

Curvularia pallescens Boedijn

Fig. 5.18

Colonies on CYA 50–65 mm diam, on MEA covering the whole Petri dish, plane, of low to floccose mycelium, pale grey to mid grey, reverse brown to dark brown. On G25N, colonies 3–6 mm diam, brown or grey. At 58C, usually germination. At 378C, colonies 40–50 mm diam, of floccose grey mycelium, sometimes brown soluble pigment, reverse dark brown. On DCMA, colonies 50–65 mm diam, plane, with pale brownish grey to mid brown mycelium, reverse reddish brown to dark brown. Conidia borne from pores in geniculate hyphae, ellipsoidal, with three septa, sometimes inconspicuous, almost straight along one side but with the eccentric swelling of the penultimate cell character-

Fig. 5.18 Curvularia pallescens (a) colony on MEA, 7 d, 258C; (b) fruiting structures in situ, bar = 50 mm; (c) conidia, bar = 10 mm

5.14 Genus Drechslera

istic of the genus, 18–25  9–12 mm, with walls smooth, pale to mid brown, and with all cells in each conidium of a similar colour. Distinctive features. This species lacks the abrupt curvature in the conidia observed in other foodborne species; conidia are relatively light in colour and characterised by an even colour in all cells. Colony reverses on CYA and MEA are brown, not blue black. Physiology. Like Curvularia lunata, C. pallescens was able to germinate at 0.86 aw, but only grew down to 0.89 aw at 258C (Hocking et al., 1994). Mycotoxins. This species is not known to produce mycotoxins. Ecology. The major known source of Curvularia pallescens is sorghum (Pitt et al., 1994; Ishrat et al., 2005). However, it has also been recorded on rice (Pitt et al., 1994; Gutierrez et al., 2002) and causing rots in melons (Sharma et al., 2002). References. Ellis (1971); Sivanesan (1987).

5.14 Genus Drechslera S. Ito Shoemaker (1959) revised the genus Drechslera, provided a clear separation from Helminthosporium, and erected Bipolaris. Leonard and Suggs (1974) segregated Exserohilum from Bipolaris. These three genera produce long, large conidia with transverse septa, which are much thicker than those in e.g. Trichoconiella. In Drechslera, conidia are cylindrical and germinate at any cell, while in Bipolaris and Exserohilum, conidia narrow gradually and germinate only from the terminal cells. Drechslera species are serious pathogens on cereal plants, and a number of species have been recorded to occur on grains from time to time. However, they appear to have little or no role in food spoilage. Species are mostly distinguished by differences in spore size; one, Drechslera triticirepentis, is treated here as an example.

Drechslera tritici-repentis (Diedicke) Shoemaker Fig. 5.19 Teleomorph: Pyrenophora tritici-repentis (Diedicke) Drechsler Colonies on CYA 55–70+ mm diam, sometimes covering the Petri dish, of low to deep, floccose,

85

pale to mid grey mycelium, sometimes lightly sporing in upper mycelial layers; exudate and soluble pigment absent; reverse dark grey to blue black. Colonies on MEA covering the Petri dish, similar to on CYA but sporulation usually absent; reverse brownish grey to bluish black. Colonies on G25N 4–8 mm diam, of sparse, pale grey mycelium. At 58C, sometimes germination. At 378C, no growth. Colonies on DCMA when grown under lights 45–70+ mm diam, similar to those on CYA, sometimes lightly sporing; reverse pale to dark grey. Conidiophores 150–250(–400) mm long, straight or sinuous, sometimes knobby terminally, bearing solitary conidia; conidia cylindroidal, commonly 70–120  14–17 mm, with five to seven septa, brown, smooth walled. Distinctive features. See genus preamble. Taxonomy. The teleomorph of this species is the Ascomycete Pyrenophora tritici-repentis (Diedicke) Drechsler. This is not usually seen in pure culture; sometimes immature perithecia may be formed (Sivanesan, 1987). Physiology. No physiological studies on Drechslera species are known to us. In common with most other plant borne dematiaceous Hyphomycetes, growth is unlikely to occur below about 0.90 aw. Species appear to be mesophilic. Mycotoxins. No mycotoxins are known to be produced. Ecology. Drechslera tritici-repentis is one cause of ‘‘pink tip’’ in wheat. We have isolated it from 56% of 344 surface disinfected North American wheat samples, where it was present at up to 27% in infected samples and 2% of all grains examined (unpublished data). Occurrence in Australian wheat was much lower (5% of samples; unpublished data) and it was not isolated from Southeast Asian commodities (Pitt et al., 1994, 1998a). Several other Drechslera species have been reported from grains and seeds. However, most records examined refer to species now included in Bipolaris or, less commonly, Exserohilum. Only D. sorokiniana (Sacc.) Shoemaker, from German durum wheat (Nirenberg et al., 1995) relates with certainty to a Drechslera species. References. Alcorn (1983); Sivanesan (1987).

86

5 Primary Keys and Miscellaneous Fungi

Fig. 5.19 Drechslera tritici-repentis (a) colony on MEA, 7 d, 258C; (b) colonies on tap water agar plus millet seed and on V-8 juice, 7 d, 258C; (c) conidia in situ, bar = 50 mm; (d) conidia, bar = 10 mm

5.15 Genus Endomyces Reess Endomyces is a genus of yeast-like Ascomycetes, indeed often classified with the yeasts, in Saccharomycopsis (Kurtzman and Fell, 1998; Barnett et al., 2000). However, Yamada et al. (1996) considered that differences in DNA from the ITS region are sufficient to consider Endomyces as a genus separate from Saccharomycopsis. As growth is filamentous

and spreading on agar plates, their decision is followed here. The conidial state consists of yeast-like cells borne on spicules or small projections. The Ascomycete state occurs sometimes in culture: the production of unenclosed, evanescent asci is suggestive of a relationship with the true yeasts. The only species common in foods is Endomyces fibuliger. See also Hyphopichia, which appears to be a closely related genus.

5.15 Genus Endomyces

87

Fig. 5.20 Endomyces fibuliger (a) colonies on CYA and MEA, 7 d, 258C; (b) conidiophores and conidia (slide culture), bar = 10 mm; (c) adherent ascospores, bar = 5 mm

Endomyces fibuliger Lindner

Fig. 5.20

Endomycopsis fibuliger (Lindner) Dekker Saccharomycopsis fibuligeria (Lindner) Klocker ¨

Colonies on CYA 18–30 mm diam and on MEA 15–25 mm diam, of low and sparse to moderately dense white to pale grey mycelium, sometimes centrally umbonate; reverse pale, off-white or very pale yellow. Colonies on G25N 8–15 mm diam, similar to those on CYA. No germination at 58C. At 378C, colonies usually 5–20 mm diam, similar to at 258C; reverse off-white to dull yellow brown. Conidia borne from spicules (small projections) along the length of undifferentiated hyphae, yeastlike, spherical, ellipsoidal or pyriform, 3–6 mm long, with thin, smooth walls. Ascospores sometimes produced in cultures on CYA or MEA, asci evanescent, ascospores observed singly or more commonly in

tight clusters of 2–4, ellipsoidal, with a longitudinal flange, offset from the spore axis to give a ‘‘bowler hat’’ appearance, 7–8 mm long, including the flange. Distinctive features. This species produces white, powdery, filamentous colonies which bear conidia from spicules on vegetative hyphae, and sometimes also by budding (yeast-like cells). Taxonomy. This species has been described under both Endomycopsis and Saccharomycopsis; in the latter case the epithet is spelled ‘‘fibuligera’’. As explained above, we consider E. fibuliger to be the valid name. More recent taxonomies list this species under Saccharomycopsis fibuligera (Lindner) Klocker (Kurtzman and Fell, 1998; Barnett et al., ¨ 2000) without taxonomic justification. This species and Hyphopichia burtonii have a close resemblance in culture. Both grow at more or less the same rates and have similar habitats. The

88

5 Primary Keys and Miscellaneous Fungi

fragmenting of fertile hyphae in H. burtonii is the only feature which provides a characteristic difference in morphology. Physiology. The physiology of Endomyces fibuliger has not been studied in any detail. A minimum growth temperature around 58C has been reported (Spicher, 1986b). E. fibuliger on wheat and rye bread was less affected by reduced O2 levels than other spoilage moulds and its growth could not be prevented by modified atmosphere packaging (Suhr and Nielsen, 2005). Mycotoxins. This type of fungus is unlikely to produce toxic compounds. Ecology. This species is not uncommon in cereals and cereal products, especially packaged bread (Spicher, 1984a, 1985). In Europe, along with Hyphopichia burtonii, it is known as ‘‘chalky mould’’ of bread (Spicher, 1986b). It produces powerful amylases (Gogoi et al., 1987; Manilal et al., 1991). Endomyces fibuliger (Saccharomycopsis fibuligera) was found in fermented potato pulp used in an Indonesian starter Ragi tape (Abe et al., 2004) and in ‘‘Marcha’’, a traditional amylolytic starter used to produce sweet-sour alcoholic drinks in the Himalayan regions of India, Nepal, Bhutan and Tibet (Tsuyoshi et al., 2005). References. Von Arx (1981b); Barnett et al. (1990; 2000).

5.16 Genus Epicoccum Link Epicoccum is a Hyphomycete genus, a saprophyte or secondary invader of senescent plant tissue. It is characterised by the production of masses of large, spherical, stalked, irregularly septate conidia, borne on rapidly growing multicoloured colonies. Epicoccum is very widely distributed in the air, in soil and on decaying vegetation, one particular source being drying grass (Kilpatrick and Chilvers, 1981). Its ubiquity in the environment means it is commonly found on foods but it is an uncommon cause of spoilage. There is a single species, Epicoccum nigrum.

Epicoccum nigrum Link

Fig. 5.21

Epicoccum purpurascens Ehrenb. ex Schltdl.

Colonies on CYA 60 mm diam or more, often covering the whole Petri dish, low and dense or funiculose or floccose; mycelium orange brown,

brown, or sometimes reddish or greenish, in fresh isolates enveloping or surmounted by brown black clusters of conidia, sometimes dominating colony appearance; clear to red brown exudate sometimes produced; reverse usually orange brown to black or with pink, red or green areas. On MEA colonies generally similar to on CYA, sometimes with a different colour combination, or occasionally with surface slimy. On G25N colonies 3–10 mm diam, low to deep, yellow to dark brown; reverse similar, sometimes with yellow soluble pigment. At 58C, response variable, no growth to colonies up to 8 mm diam formed. No growth at 378C. Conidia borne solitarily on short conidiophores, usually in dense clusters, spherical with a broad, tapering, truncate base; brown, irregularly septate when mature, commonly 15–25 (–30) mm diam, with rough walls obscuring numerous septa. According to Schol-Schwarz (1959), conidia on stems of sterile lupins measure 7–65  6–54 mm. As has been observed with other genera, conidia on agar media are much less variable in size and often smaller than on natural substrates. Distinctive features. See the genus preamble. Taxonomy. Kilpatrick and Chilvers (1981) examined the variability of 2000 isolates of Epicoccum and concluded that all belonged to a single, genetically variable species. A study of the 5.8S and ITS regions of DNA confirmed this (Wang and Guo, 2004). In early literature this species was usually known as E. nigrum Link. With the gradual acceptance of the 1821 starting date (See Chapter 3), this name was in time replaced by E. purpurascens. Now, with the reversion to the 1753 starting date, E. nigrum is again the correct name. Physiology. According to Kilpatrick and Chilvers (1981), maximum growth rates vary widely among isolates of this species. However, optimal temperatures for growth are usually 20–258C, with a maximum at 30– 358C and a minimum below 58C. The optimum water potential for growth is –20 bars (0.98 aw) and minimum –120 bars (0.91 aw) (Kilpatrick and Chilvers, 1981). Mycotoxins. Mycotoxin production has not been recorded. Ecology. As noted in the genus preamble, Epicoccum is common in the general environment and hence readily finds its way onto foods such as cereals and nuts. It has been reported to cause spoilage of cantaloupes, in which it produces a

5.17 Genus Fusarium

89

Fig. 5.21 Epicoccum nigrum (a) colonies on CYA and MEA, 7 d, 258C; (b) conidiophores and conidia, bar ¼ 10 mm; (c) conidia, bar ¼ 10 mm

red discolouration (Snowdon, 1991), and as a pathogen on cucumbers, tomatoes, apples and pears (Bruton et al., 1993). Among other fungi, E. nigrum can produce core rot of apples (Combrink et al., 1985). It has been associated with spoilage of pecans and is sometimes present in barley after harvest and during malting, but is not a cause of gushing (see Pitt and Hocking, 1997; Niessen et al., 1991; Hudec, 2007). E. nigrum has been reported on fresh vegetables, nuts and cereals. Other records include rice, wheat, maize and wheat; gourds and muskmelons, pecans, peanuts, frozen and cured meats and biltong (see Pitt and Hocking, 1997; Gros et al., 2003).

Epicoccum nigrum was isolated from 29% of soybean samples and 7% of cashew samples in Thailand, with up to 12% infection in infected soybean samples, 15% in cashews and 1% infection overall in both (Pitt et al., 1994). References. Schol-Schwarz (1959); Domsch et al. (1980); Kilpatrick and Chilvers (1981).

5.17 Genus Fusarium Link The character which defines the genus Fusarium is the production of septate, fusiform to crescentshaped conidia, termed macroconidia, with a foot-

90

shaped basal cell and a more or less beaked apical cell. Macroconidia may be produced in discrete pustules, called sporodochia, or in confluent, slimy masses, known as pionnotes. Mounts from these areas, which are usually cream, salmon pink or orange, will reveal masses of these spores. However, they are rarely widely distributed over the colony and frequently are entirely absent. A light bank is invaluable for promoting formation of macroconidia in Fusarium cultures. Many species of Fusarium also produce smaller one to two celled conidia, microconidia, of various shapes. Chlamydoconidia, either terminal or intercalary, are characteristic of some species also. Fusarium colonies are usually fast growing and consist of felty aerial mycelium which may be pale, or brightly coloured in shades of pink, red, violet or brown. Fusarium species are renowned for their role as plant pathogens, causing a wide range of diseases such as vascular wilts, root and stem rots, pre- and post-emergence blight and many others. Leslie and Summerell (2006) include 70 species in their laboratory manual but the taxonomy of Fusarium is still undergoing change, with biological and phylogenetic criteria as well as morphological concepts used to delineate species (Leslie and Summerell, 2006). Fusarium species are widely distributed in soils, particularly cultivated soils, and are active in the decomposition of cellulosic plant materials. They are a major cause of storage rots of fruits and vegetables and are commonly associated with cereals and pulses, which they usually invade before harvest. De Nijs et al. (1996) have published a compendium of the occurrence of more than 60 Fusarium species in raw materials, foods and feeds but, in our experience, a relatively small number of species are important in food spoilage. A number of Fusarium species have teleomorphs belonging to the genera Nectria Fr., Calonectria De Not. and Gibberella Sacc. These teleomorphs are perithecial Ascomycetes, and are generally not produced in culture. However, fertile strains of F. solani sometimes form reddish brown perithecia on PDA and F. graminearum usually produces dark purple perithecia on Carnation Leaf Agar. These teleomorphs are known as Nectria haematococca Berk. and Broome and Gibberella zeae (Schwein.) Petch, respectively (Booth, 1971).

5 Primary Keys and Miscellaneous Fungi

Taxonomy. A bewildering array of taxonomic systems have been applied to Fusarium, ranging from the milestone work of Wollenweber and Reinking (1935), which distinguished 65 species, to the simplification by Snyder and Hansen, who accepted only 9 species in a series of papers in the 1940s. The interrelationships of the principal taxonomic schemes for Fusarium are discussed by Booth (1971), Nelson et al. (1983) and more recently by Leslie and Summerell (2006). In recent years, Fusarium taxonomy and nomenclature has undergone many changes, with molecular methods elucidating the existence of cryptic species within some of the broader species concepts. In addition, many new species have been described, often from natural ecosystems rather that economically important crops (see Leslie and Summerell, 2006). The nomenclature of Fusarium species described in this work follows that of Leslie and Summerell (2006) which is based on Nelson et al. (1983) which, in turn, is based on Wollenweber and Reinking (1935). Reference is made to the names used by Nelson et al. (1983) where they differ from those adopted by Leslie and Summerell (2006). Mycotoxins. Fusarium is one of the three major fungal genera producing toxins. The most widespread Fusarium mycotoxins are the trichothecenes, a family of sesquiterpenes. More than 50 such compounds are known to be produced by species in this genus. Some are highly toxic: none appears to be benign. Trichothecenes are often produced as mixtures even under pure culture conditions and are very difficult to separate, so the toxicity of many compounds remains uncertain. The most important are deoxynivalenol (and sometimes nivalenol), produced by F. graminearum, F. culmorum and related species. Zearalenone, not a true mycotoxin, really an oestrogen analogue produced by fungi, is formed by the same species that make deoxynivalenol. T-2 toxin, produced primarily by Fusarium sporotrichioides, caused the deaths of many people and animals during the 20th century, but is of uncommon occurrence now. Some Fusarium species produce other types of toxic compounds. The most important of these are the fumonisins, produced by F. verticillioides, F. proliferatum and several other species. Fumonisins are sphingosine analogues, interfering with membrane function in man and animals (Miller and Trenholm, 1994). Other

5.17 Genus Fusarium

less toxic or less important compounds, such as moniliformin and fusaric acid, are discussed briefly under the appropriate species. Deoxynivalenol and nivalenol. Like all trichothecene mycotoxins, deoxynivalenol (DON) and nivalenol are inhibitors of protein synthesis (Feinberg and MacLaughlin, 1989). Nivalenol is much more toxic than DON, but is produced in much lower quantities in grains, and is considered a less significant mycotoxin (Miller et al., 2001). DON is especially toxic to pigs, where, at quite low intake levels (< 5 mg/kg in feed) it causes vomiting, feed refusal and reduced weight gain due to neurotoxic effects. Cattle and poultry are resistant to reasonable levels (> 5 mg/kg in feed) of DON (Miller et al., 2001). In humans, high doses of DON may cause abdominal pains, dizziness, headache, nausea, vomiting and other effects. Cases of such acute poisoning are rare, but have been recorded from India (Bhat et al., 1989), Japan (Udagawa, 1988) and several outbreaks in China (JECFA, 2001). DON is also immunosuppressive, and because of its widespread occurrence in grains, is undoubtedly responsible for susceptibility to bacterial and viral diseases in man and animals (Miller et al., 2001). The most comprehensive study of DON including occurrence, toxicity, analytical methods and intake levels is that by JECFA (2001). They concluded that DON is not significantly carcinogenic, mutagenic or teratogenic (JECFA, 2001). The Committee set a provisional tolerable daily intake of 1 mg/kg weight per day. They were unable to estimate the level of DON in foods below which any acute effect in humans would occur. Zearalenone. Zearalonone is one of the five most significant mycotoxins (Miller, 1995). It is not a true mycotoxin, being a nonsteroidal oestrogen produced by a fungus. It is not acutely toxic and has not been associated with any fatal disorder in animals or humans. However, it has caused oestrogenic syndromes in pigs, and perhaps in human adolescents as well. Zearlaenone is produced by the same Fusarium species that produce deoxynivalenol, i.e. F. graminearum and related species, especially F. culmorum, F. equiseti and F. crookwellense. The association of these fungi with cereal crops is worldwide, as is the production of zearalenone (Marasas et al., 1984).

91

Pigs are especially sensitive to zearalenone. Signs of hyperoestrogenism generally appear at dietary levels > 1 mg/kg, but can occur at lower levels sometimes. In prepubertal gilts, clinical signs include vulval swelling, uterine enlargement and mammary development. Mature sows can develop ovarian atrophy, constant oestrus and pseudopregnancy (Hagler et al., 2001). Prepubertal male pigs can undergo a feminising effect with mammary development, decreased testicular size and loss of libido, but mature boars are resistant (Hagler et al., 2001). It was once claimed that humans had shown similar signs in Puerto Rico, but an FDA investigation failed to confirm this (Goodman et al., 1987). T-2 toxin. T-2 toxin is the cause of alimentary toxic aleukia (ATA) a devastating disease which occurred in the USSR during and after Second World War, in times of extreme food shortage resulting in consumption of overwintered cereals (Joffe, 1978; Beardall and Miller, 1994). Many people, probably hundreds of thousands, died as a result (Marasas et al., 1984). ATA was characterised by leucopoenia, bleeding from nose, throat and gums, haemorrhagic rash, exhaustion of the bone marrow and fever. Vomiting, nausea, diarrhoea and abdominal pain also usually occurred. Decrease in immunological functions led to susceptibility to bacterial and viral diseases, and often death (Joffe, 1978; Beardall and Miller, 1994). Haemorrhagic syndrome in cattle, pigs and poultry in the United States in the 1960s was also probably due to T-2 toxin (Desjardins, 2006). T-2 toxin is produced by F. sporotrichioides and, less commonly, F. poae (Desjardins, 2006). It appears to be produced only under cold conditions, and fortunately is now uncommon. The occurrence, toxicity and biology of T-2 toxin were examined in detail by JECFA (2001). They established that T-2 (and its metabolite HT-2) were immunotoxic and haemotoxic compounds in several animal species after short-term intake. Long-term effects could not be evaluated. T-2 was at most weakly genotoxic. In the absence of long-term studies, T-2 was not classifiable as to carcinogenicity (JECFA, 2001). Fumonisins. Fumonisins are produced by F. verticillioides (formerly known as F. moniliforme), by the closely related species F. proliferatum, uncommonly by F. subglutinans and F. oxysporum,

92

and about 10 other species not treated here. The first two named species are by far the most important. F. proliferatum is one cause of maize ear rot in Europe and is a pathogen on a variety of other crops. F. verticillioides is endemic in maize and found wherever that crop is grown. This species grows well at higher temperatures, and ear rot and fumonisin accumulation are associated with drought, insect stress and growing hybrids outside their areas of adaptation (Miller, 2001, 2008). Fumonisins are analogues of sphingosines, essential components of cell membranes. Fumonisins have been reported to interfere specifically with dihydroceramide, an essential enzyme in the biosynthesis of sphingosine and related compounds (Wang et al., 1991). Fumonisins have been found to be biologically active in many animal species, but horses and pigs are the most affected. In horses, fumonisins are responsible for leucoencephalomalacia, a rapidly progressing disease that causes tissue liquefaction in equine brains. It has been known in the United States for more than 100 years that feeding maize to horses is a dangerous practice (Desjardins, 2006): it was not until 1988 that the reason became apparent (Gelderblom et al., 1988; Marasas et al., 1988). For horses, consumption of feed containing > 10 mg/kg fumonisin B1 in the diet (equivalent to 0.2 mg/kg body weight per day) was associated with increased risk of developing this disease (JECFA, 2001). In pigs, fumonisins cause pulmonary oedema, due to left ventricle heart failure (JECFA, 2001), while in rats the primary effect is to cause liver cancer (Gelderblom et al., 1991), but they also cause programmed cell death (apoptosis; Tolleson et al., 1996). It is unusual for a single toxin to have such diverse effects in different animal species, and the reasons for that remain unknown (Desjardins, 2006). In humans, fumonisins and Fusarium verticillioides are associated with oesophageal cancer. Extensive studies in areas of low and high maize consumption in South Africa have established this connection (Rheeder et al., 1992; JECFA, 2001). This disease is also prevalent in areas of China and occurs at significantly higher levels than background also in parts of Iran, northern Italy, Kenya and a small area of the southern USA (JECFA, 2001). In all of those areas consumption of maize and maize products is very high (JECFA, 2001). There is also increasing evidence that fumonisins are associated

5 Primary Keys and Miscellaneous Fungi

with neural tube defects such as spinal bifida in a population along the Texas–Mexican border (Desjardins, 2006). For the human population, a provisional maximum tolerable daily intake (PMTDI) of 2 mg/kg body weight per day was established by JECFA (2001). Determining which particular Fusarium species produces which mycotoxins has not been easy. Unstable taxonomy and misidentification have combined to cloud the picture. In a major contribution, Marasas et al. (1984) examined several hundred Fusarium isolates, identifying them according to Nelson et al. (1983) and then assessing mycotoxin production. Sections on mycotoxins in the species descriptions which follow are based on Marasas et al. (1984), Desjardins (2006) and Leslie and Summerell (2006). Claims concerning mycotoxin production by particular species in more recent references should be treated with some caution. Cultural instability. Many Fusarium species are notorious for their instability in culture. Isolates of some species will degenerate quickly, often after only one or two transfers. For this reason, it is important to identify Fusaria as soon as possible after primary isolation. Pure cultures for identification are traditionally started from a single germinated spore, as the mass transfer of Fusaria appears to increase the rate of deterioration of strains in culture. Identification procedures. Identification of Fusarium isolates is often difficult, but the task can be made easier by observing a few basic rules:

 identify cultures as soon as possible after primary isolation;

 always grow cultures for identification from single germinated conidia; standardised media and incubation conditions;  use a light bank (Chapter 4) whenever possible.

 use

Diagnostic features. The main characters used to distinguish species of Fusarium are (1) the size and shape of the macroconidia; (2) the presence or absence of microconidia; (3) the manner in which microconidia are produced; (4) the type of phialide on which microconidia are produced; (5) the presence or absence of chlamydoconidia; and (6) the colours and morphology of colonies on PDA.

5.17 Genus Fusarium

93

The morphology of macroconidia is a principal diagnostic feature for Fusarium species. Macroconidia generally have at least three septa, with a differentiated apical cell which may be pointed, rounded, hooked or filamentous, and a basal cell which may be foot-shaped, with a distinct heel, or just slightly notched. Fusarium macroconidia generally exhibit some degree of curvature, the convex and concave sides being referred to as the dorsal and ventral sides, respectively. Although some macroconidia are usually produced in the aerial mycelium, the shape and size of those in sporodochia are more regular and are used for identification purposes where possible. Microconidia are usually produced in the aerial mycelium and their shape can be very important in Fusarium identification. Most species which produce microconidia form only a single type, the most common shape being ellipsoidal to clavate. However, F. poae produces spherical to apiculate microconidia and F. sporotrichioides produces a variety of shapes: ellipsoidal, pyriform and spherical. The method of production of microconidia and the types of phialides on which they are borne are also useful diagnostic criteria. F. verticillioides produces its microconidia in long, delicate, dry chains, which are best observed by using the 10 objective of the

compound microscope. Some species produce microconidia in false heads (small, mucoid, adherent balls of conidia) and others produce them singly. In some species, microconidia are produced on phialides with only one pore, and these are termed monophialides, but a few species produce phialides with more than one pore (polyphialides). Species which produce polyphialides usually produce monophialides as well. Descriptions. Descriptions of the microscopic features of species in this book are based on structures formed in cultures grown on DCPA at 258C, with a 12 h photoperiod, under a light bank consisting of two cool white fluorescent tubes and one black light tube (see Chapter 4). Cultures should be examined at 7 days, then at 10 days and 14 days if sporulation is poor. Aerial mycelium often develops better after 10 days incubation on DCPA and chlamydoconidium production is more reliable in older cultures. Descriptions of the colony characteristics are taken from cultures grown on PDA at 258C for 7 days, also with a 12 h photoperiod. Additional information may be gained by recording the growth rates of colonies on PDA at 258C and 308C after 3 days (Burgess et al., 1994). Growth rates at 308C on PDA are particularly helpful in distinguishing isolates of Fusarium avenaceum from those of Fusarium acuminatum.

Key to Fusarium species included here 1

Microconidia abundant Microconidia rare or absent

2 9

2 (1)

Colonies on PDA with mycelium and/or reverse coloured greyish rose or burgundy Colonies on PDA in shades of cream, pale salmon or violet

3 5

3 (2)

4 (3)

5 (2)

6 (5)

Microconidia spherical to apiculate, borne singly on monophialides Microconidia ellipsoidal, clavate, fusiform and/or pyriform, borne on polyphialides or both polyphialides and monophialides Microconidia clavate only, produced profusely, giving colonies on PDA a powdery appearance Microconidia various shapes: clavate, pyriform and spindle shaped (check PDA cultures also) Microconidia produced in long or short chains (some false heads may also be present) Microconidia produced singly or in false heads Microconidia produced from monophialides only Monophialides and polyphialides both present

F. poae 4

F. chlamydosporum F. sporotrichioides

6 7 F. verticillioides F. proliferatum

94

5 Primary Keys and Miscellaneous Fungi

7 (5)

Colonies cream or bluish, sporodochia cream Colonies pale salmon or violet, sporodochia salmon

8 (7)

Microconidia borne on short, stout monophialides; chlamydoconidia usually produced Microconidia borne on polyphialides and slender monophialides; chlamydoconidia not produced

9 (1)

Colonies cream, pale salmon or brown Colonies greyish rose to burgundy

10 (9)

Macroconidia cigar- or spindle-shaped, produced in the aerial mycelium Macroconidia obviously curved, produced in sporodochia

11 (9)

Macroconidia robust, ventral side straight; aerial mycelium tan to brown Macroconidia delicate and slender, slightly or definitely curved; aerial mycelium white or pinkish

12 (11)

Macroconidia longer and narrower, maximum width 5.5 mm 13 (11)

Macroconidia with elongated basal cell and long, whip-like apical cell Macroconidia with basal and apical cells not obviously elongated

14 (13)

Macroconidia delicate and needle-like, with sides almost parallel Macroconidia with slight to definite curvature

Telemorph: Gibberella acuminata Wollenw. Colonies on CYA 40–50 mm diam, of dense, felty mycelium, white to greyish rose or greyish magenta; reverse uniformly pale or with areas of greyish rose. Colonies on MEA 45–65 mm diam, yellow brown centrally, greyish rose at the margins; reverse deep brownish yellow to brownish orange, occasionally pale. Colonies on G25N 9–15 mm diam. At 58C, colonies 7–12 mm diam. No growth at 378C. On PDA, colonies usually covering the whole Petri dish, of dense to floccose white to pale salmon mycelium, sometimes greyish rose at the margins; reverse dark ruby centrally, greyish ruby at the margins. On DCPA, colonies sparse, of floccose to funiculose white to pale salmon mycelium; reverse pale or with brownish red annular rings. Macroconidia relatively slender, usually with five septa, but three and four septa not uncommon, with a long, tapering apical cell and foot-shaped basal cell, distinctly but not highly curved, with

F. oxysporum F. subglutinans 10 11

Macroconidia short and stout, up to 7 mm wide

Fusarium acuminatum Ellis & Everh. Fig. 5.22

F. solani 8

F. semitectum F. equiseti 12 13 F. culmorum (See F. graminearum) F. graminearum F. longipes 14 F. avenaceum F. acuminatum

the widest point often one third of the distance from the base, giving a ‘‘bottom-heavy’’ appearance; microconidia produced sparsely by some isolates; chlamydoconidia produced, relatively slowly. Distinctive features. Ruby to dark ruby reverse colours on PDA, and relatively slender, slightly curved macroconidia, usually with five septa, are the distinctive features of Fusarium acuminatum. However, unless chlamydoconidia are present, this species can be confused with F. avenaceum (see below). F. armeniacum is very closely related (Burgess and Summerell, 2000). Taxonomy. Perithecia of Gibberella acuminata Wollenw., the teleomorph of F. acuminatum, are formed in the laboratory when opposite mating types are inoculated onto sterile wheat straws (Booth, 1971). Isolates of F. acuminatum show considerable variability in culture, and this variability, correlated with secondary metabolite production (Logrieco et al., 1992), has been shown to be due to the fact that two species were included within F. acuminatum. F. armeniacum (Forbes et al.) L.W. Burgess and Summerell, previously described as F. acuminatum var. armeniacum G.A.

5.17 Genus Fusarium

95

Fig. 5.22 Fusarium acuminatum (a) colonies on PDA and DCPA, 7 d, 258C; (b, c) macroconidia, bar = 10 mm

Forbes et al. has been elevated to species level (Burgess and Summerell, 2000) being distinguished on morphological grounds, isoenzyme patterns, molecular markers and mycotoxin profiles. Macroconidia of F. armeniacum are produced in distinct apricot coloured sporodochia. Physiology. Some isolates of Fusarium acuminatum have antioxidant enzyme activity (Kayali and Tarhan, 2005). Mycotoxins. Most of the mycotoxin production reported from Fusarium acuminatum, particularly the production of trichothecene toxins, is probably more correctly due to F. armeniacum (Desjardins, 2006; Leslie and Summerell, 2006). Isolates producing type A trichothecenes including T-2 toxin, HT-2 toxin and diacetoxyscirpenol (see Pitt and Hocking, 1997) are most likely to be F. armeniacum. In a survey of 25 isolates of F. acuminatum from different sources and geographic locations,

Logrieco et al. (1992) divided the isolates into three categories, (a) enniatin B and/or moniliformin producers (probably F. acuminatum sensu stricto), (b) T-2, HT-2 and/or neosolaniol producers (probably F. armeniacum) and (c) nontoxigenic. F. acuminatum sensu stricto produces moniliformin (Chelkowski et al., 1990; Logrieco et al., 1992) and enniatins (Logrieco et al., 1992; Kononeko et al., 1993; Desjardins, 2006; Leslie and Summerell, 2006) as well as some other minor toxins (Desjardins, 2006). Ecology. Fusarium acuminatum has been isolated from a wide variety of plants throughout the world. Although some isolates may cause severe root rot in particular legume species (Leslie and Summerell, 2006), F. acuminatum is generally regarded as a saprophyte. It has been reported to cause rot in pumpkins (Elmer, 1996), is one cause of rot in stored potatoes (Theron and Holz, 1990) and is weakly pathogenic in

96

5 Primary Keys and Miscellaneous Fungi

bananas (Jime´nez et al., 1993). It is quite common in poor quality wheat from cool temperate zones (Mills and Wallace, 1979; Abramson et al., 1987). It has been isolated from developing peanut pods (Barnes, 1971), barley (Abdel-Kader et al., 1979) and, in our laboratory, from rain damaged sorghum and soybeans. The incidence of F. acuminatum in tropical commodities was low (Pitt et al., 1993, 1994). References. Domsch et al. (1980), under G. acuminata; Nelson et al. (1983); Desjardins (2006); Leslie and Summerell (2006).

Fusarium avenaceum (Fr.) Sacc.

Fig. 5.23

Teleomorph: Gibberella avenacea R.J. Cooke Colonies on CYA covering the whole Petri dish, moderately deep to deep, of open, floccose

mycelium coloured white, very pale rose or deeper greyish rose; reverse varying from pale to pale yellow or with areas of greyish rose or sometimes uniformly deep burgundy. Colonies on MEA 45– 55 mm diam, low to moderately deep, of open floccose to funiculose mycelium, coloured white, pale rose or greyish rose, sometimes brown centrally; reverse brownish orange, sometimes paler centrally or at the margins. Colonies on G25N 9– 15 mm diam. At 58C, colonies 10–12 mm diam. No growth at 378C. On PDA, colonies moderately deep to deep, of dense mycelium coloured white, pale salmon or sometimes dark brownish red, with central masses of reddish orange sporodochia, sometimes surrounded by an outer ring of paler sporodochia; reverse greyish red, with darker annular rings, paler towards the margins. On DCPA, colonies deep, of moderately dense white to pale salmon mycelium with a central mass of orange to

Fig. 5.23 Fusarium avenaceum (a) colonies on PDA and DCPA, 7 d, 258C; (b, c) macroconidia, bar = 10 mm

5.17 Genus Fusarium

salmon sporodochia, often surrounded by concentric rings of sporodochia; reverse pale. Macroconidia long, slender, with four to seven septa, thin-walled, straight or slightly curved, with a tapering apical cell and a notched or foot-shaped basal cell; microconidia produced sparsely by some isolates; chlamydoconidia absent. Distinctive features. Fusarium avenaceum is distinguished by thin walled, needle-like macroconidia and by the absence of chlamydoconidia. Despite the fact that F. avenaceum and F. acuminatum are not considered by Fusarium taxonomists to be closely related, these two species can be difficult to distinguish, as isolates with macroconidia of intermediate form are not uncommon. Colony diameters on PDA at 308C after 3 days can be a useful differentiating feature: under these conditions colonies of F. avenaceum are usually 8– 15 mm diam, whereas those of F. acuminatum are 15–28 mm diam (Burgess et al., 1994). Isolates of F. avenaceum show an unusually broad range of colours on PDA and also have a very broad host range. However, extensive genetic analysis has shown no bases from splitting the species, and pathogenicity tests on single strains have confirmed the broad host range (Nalim, 2004). Taxonomy. The teleomorph of Fusarium avenaceum is Gibberella avenacea R.J. Cook. It is not usually seen in culture on the media used here. Physiology. The optimum growth temperature for Fusarium avenaceum is 358C, the minimum near –38C and the maximum 318C (Domsch et al., 1980). The minimum aw for growth is approximately 0.90 at 258C (Magan and Lacey, 1984c), and the pH optimum ranges between 5.4 and 6.7 (Domsch et al., 1980). Mycotoxins. This species has been reported to produce a variety of trichothecene and other mycotoxins. However, Nelson et al. (1983) regarded only reports of moniliformin production as accurate. Later reports have confirmed this (Chelkowski et al., 1990; Abbas et al., 1991; Bosch and Mirocha, 1992). Reports of production of fusarin C (Farber and Sanders, 1986; Thrane, 1988; Leonov et al., 1993) and enniatins (Blais et al., 1992; Kononeko et al., 1993) also appear to be reliable (Desjardins, 2006; Leslie and Summerell, 2006). Production of any trichothecene toxin has not been confirmed, and Fusarium avenaceum does not carry the tri5 gene which is essential for trichothecene production (Tan and Niessen, 2003).

97

Ecology. Fusarium avenaceum has a worldwide distribution wherever crops are grown, but is relatively uncommon in food commodities. It is a major component of Fusarium head blight in cereals in Europe, the US Pacific Northwest region and Canada (Desjardins, 2006). Logrieco et al. (2002) identified F. avenaceum as a component of Fusarium ear rot of maize in Europe. It has been reported from barley (Flannigan, 1969; Petters et al., 1988; Stenwig and Liven, 1988) where it may inhibit germination of malting grains (Hudec, 2007), but is of minor importance in gushing of beer (Niessen et al., 1992). Other reported sources are sorghum (Onyike and Nelson, 1992), peanuts (Joffe, 1969), pigeon peas (Maximay et al., 1992) and, in our laboratory, triticale. F. avenaceum has been reported to cause spoilage of cool-stored broccoli (Mercier et al., 1991), dry rot of stored carrots in Italy (Marziano et al., 1992) and dry rot of rutabaga (swede turnip) in Canada (Peters et al., 2007). It has occasionally caused spoilage of apples, pears, asparagus, tomatoes, eggplant and potatoes (Snowdon, 1990, 1991) and has been reported as a postharvest pathogen of stonefruit in New Zealand (Hartill and Broadhurst, 1989). References. Domsch et al. (1980), as G. avenacea; Nelson et al. (1983); Leslie and Summerell (2006).

Fusarium chlamydosporum Wollenw. & Reinking Fig. 5.24 Fusarium fusarioides (Gonz. Frag. & Cif.) Booth

Colonies on CYA covering the whole Petri dish, of low to moderately deep floccose mycelium, coloured white to pale rosy pink, often with surface appearing powdery due to production of microconidia; reverse pale to greyish rose or brownish red. Colonies on MEA 55–70 mm diam, of low, moderately dense mycelium in shades of yellow brown, or greyish rose to greyish ruby, paler at the margins; reverse deep yellow brown to orange brown. Colonies on G25N 15–20 mm diam. At 58C, colonies 1–2 mm diam. At 378C, colonies 5–15 mm diam. On PDA, colonies of felty mycelium, coloured pale salmon, sometimes browner, or with patches of greyish red, often with a powdery appearance from profuse microconidial production; reverse deep violet brown to dark ruby, paler at the margins. On DCPA, colonies of sparse, floccose, pale salmon mycelium, often powdery with microconidia, showing poorly defined annulations; macroconidia

98

5 Primary Keys and Miscellaneous Fungi

Fig. 5.24 Fusarium chlamydosporum (a) colonies on PDA and DCPA, 7 d, 258C; (b, c) polyphialides, bar = 10 mm; (d) macroconidia, bar = 10 mm; (e) microconidia, bar = 10 mm

occasionally produced near the colony centres in salmon sporodochia; reverse pale. Macroconidia often rare, relatively short and stout, usually with three to five septa, slightly curved; microconidia produced abundantly from polyphialides in the aerial mycelium, with zero to two septa, fusiform to slightly clavate. Chlamydoconidia usually abundant in older cultures, produced singly, in pairs or in clumps. Distinctive features. The presence of abundant fusiform microconidia borne on polyphialides is the most outstanding feature of Fusarium chlamydosporum. Also colonies on PDA have dark violet brown to dark ruby reverse colours.

Taxonomy. Fusarium chlamydosporum has priority over F. fusarioides as the correct name for this species (Domsch et al., 1980; Nelson et al., 1983; Leslie and Summerell, 2006). Physiology. This species has minimum, optimum and maximum temperatures for growth of 5, 27 and 378C (Seemu¨ller, 1968). Mycotoxins. Production of type A trichothecenes (including T-2 toxin, HT-2 toxin, monoacetoxyscirpenol, neosolaniol and iso-neosolaniol) by Fusarium chlamydosporum was reported by Park and Chu (1993); however, subsequent studies have found no evidence of trichothecene production in this species (Desjardins, 2006). Moniliformin is the

5.17 Genus Fusarium

major mycotoxin produced by F. chlamydosporum (Marasas et al., 1984; Desjardins, 2006). Ecology. Fusarium chlamydosporum is mainly an inhabitant of soils in warmer climates (Domsch et al., 1980; Leslie and Summerell, 2006), and is not regarded as a plant pathogen or spoilage fungus. However, it is commonly isolated from grains in drier areas, particularly in the Middle East, southern Europe, central Asia and Australia (Leslie and Summerell, 2006), and has also been isolated from pearl millet (Wilson et al., 1993; Jurjevic et al., 2007), pecans (Huang and Hanlin, 1975) and sorghum (Rabie et al., 1975; Onyike and Nelson, 1992). A low incidence of F. chlamydosporum was found in peanuts from both Indonesia and the Philippines (Pitt et al., 1998a) and from mung beans and sorghum in Thailand (Pitt et al., 1994). Involvement in dry rot of potatoes has also been reported (Somani, 2004; Esfahani, 2006). References. Domsch et al. (1980); Nelson et al. (1983); Leslie and Summerell (2006).

99

Fusarium culmorum (W.G. Smith) Sacc. Fig. 5.25 Colonies on CYA covering the whole Petri dish, of dense felty mycelium, often with a floccose overlay, sometimes reaching the Petri dish rim, pale red to pastel red; reverse pastel red to deep red. Colonies on MEA 60 mm or more diam, floccose, in age often reaching the Petri dish lid, pale red to pastel red, commonly with a greyish orange to yellowish brown overlay; reverse brown to reddish brown. Colonies on G25N usually 5–10 mm diam, mycelium orange white, reverse yellow to orange. At 58C, germination. No growth at 378C. On PDA, colonies covering the whole Petri dish, of dense to floccose mycelium, pale red and pale yellow brown; reverse red to deep red. On DCPA, colonies 50–65 mm diam, of sparse mycelium, orange to pinkish white, bearing abundant macroconidia in orange sporodochia; reverse dull orange brown.

Fig. 5.25 Fusarium culmorum (a) colonies on PDA and DCPA, 7 d, 258C; (b, c) macroconidia, bar = 10 mm

100

Macroconidia relatively short, wide and only slightly curved, with four to five septa, 30–45 mm long, with rounded or sometimes papillate apical cells; basal cells with a slight to definite notch, sometimes papillate. Microconidia not produced. Chlamydoconidia sometimes formed, in conidia, or intercalary in the hyphae, singly or in chains, 9–14 mm diam, smooth walled. Distinctive features. Short, stout macroconidia are the prime feature distinguishing Fusarium culmorum from most other species. F. culmorum may be confused with F. crookwellense L.W. Burgess et al., but macroconidia of the latter species have a distinctly foot-shaped basal cell, whereas those of F. culmorum are shorter and stouter, and the basal cell is not distinctly foot-shaped. Taxonomy. No teleomorph is known for this species. Physiology. Fusarium culmorum has been reported to be psychrotrophic, growing down to 08C, with an optimum at 218C and a maximum of only 318C (Arsvoll, 1975); however, Magan and Lacey (1984c) reported growth at 358C. The minimum aw for growth is 0.87 at 20–258C and pH 6.5: at pH 4.0, growth did not occur below 0.90 aw (Magan and Lacey, 1984a). A strain of F. culmorum produced deoxynivalenol optimally at 258C, but only between 0.995 and 0.97–0.96 aw (Hope and Magan, 2003; Hope et al., 2005). At 158C, deoxynivalenol was produced in lower concentrations later in the growth cycle, but over a slightly greater aw range (0.995 to 0.95–0.94 aw). The dynamics of nivalenol production for this strain was similar (Hope and Magan, 2003). Zearalenone production by F. culmorum was reported to be optimal above 258C (Bottalico et al., 1982). F. culmorum is very tolerant of low O2 tensions (Magan and Lacey, 1984b). Radiation resistance of F. culmorum was relatively high: up to 0.8 kGy were needed for a tenfold reduction in spore numbers on grain, and up to 1.39 kGy on media (O’Neill et al., 1991). Mycotoxins. Fusarium culmorum produces a variety of mycotoxins: indeed the list in our files is of more than 40 compounds. However, the most important toxins confirmed to be produced by this species (Nelson et al., 1983; Marasas et al., 1984; Desjardins, 2006; Leslie and Summerell, 2006) are deoxynivalenol, nivalenol and their derivatives (Abramson et al., 2001; Hestbjerg et al., 2002;

5 Primary Keys and Miscellaneous Fungi

Chandler et al., 2003; Jennings et al., 2004) and zearalenone (Bakan et al., 2001; Hestbjerg et al., 2002; Llorens et al., 2004a; Brinkmeyer et al., 2005). Moniliformin production was reported by Scott et al. (1987) but was not detected in 42 isolates of F. culmorum from Canada by Abramson et al. (2001). Reports of production of type A trichothecenes (T-2 toxin, HT-2 toxin) have not been substantiated (Leslie and Summerell, 2006). The existence of two chemotypes of Fusarium culmorum, those that produce deoxynivalenol and those that produce nivalenol (Miller et al., 1991), has been confirmed by molecular studies. Within the trichothecene gene cluster, isolates possessing the Tri7 and Tri13 genes produce nivalenol and related compounds, whereas sequences in the Tri3, Tri5 and Tri6 genes are associated with deoxynivalenol production (Chandler et al., 2003; Jennings et al., 2004; Quarta et al., 2005, 2006). Of 55 European isolates examined by Quarta et al. (2005), 11 were the nivalenol chemotype, and the remainder were deoxynivalenol producers. Jennings et al. (2004) examined 153 isolates from England and Wales, and found that the DON chemotype was dominant over the NIV chemotype (59% vs 41%, respectively). Lauren et al. (1992) examined 45 isolates of F. culmorum from New Zealand soil and pasture and found none produced deoxynivalenol or its monoacetyl isomers. Two chemotypes were identified, one producing diacetylnivalenol with culmorin as the major metabolite accounted for 95% of isolates, while the other chemotype produced diacetoxyscirpenol. Ecology. This species has a worldwide distribution in soil and as a pathogen of cereals and other hosts, with a higher incidence in temperate climates (Domsch et al., 1980; Nelson et al., 1983; Leslie and Summerell, 2006). It is an important component of the cohort of Fusarium species that cause head blight of wheat and associated cereal crops in Europe, Canada, China and other areas with cool weather during the growing season (Desjardins, 2006). In wheat it causes extensive internal damage to the grain, and reductions in flour yield and baking quality (Meyer et al., 1986). F. culmorum was reported as the dominant Fusarium species on barley in Europe (see Pitt and Hocking, 1997). It also occurs in triticale (Perkowski et al., 1988). F. culmorum has been identified as a component of Fusarium ear rot of maize in Europe (Logrieco

5.17 Genus Fusarium

101

et al., 2002). F. culmorum was one cause of crown rot in bananas (Wade et al., 1993), and is a minor cause of spoilage of apples and pears (Snowdon, 1990). References. Domsch et al. (1980); Nelson et al. (1983); Desjardins (2006); Leslie and Summerell (2006).

Fusarium equiseti (Corda) Sacc.

Fig. 5.26

Teleomorph: Gibberella intricans Wollenw. Colonies on CYA filling the whole Petri dish, often to the lid, of dense to floccose white mycelium; reverse pale or pale salmon. Colonies on MEA

covering the whole Petri dish, of open, floccose white to pale brown mycelium; reverse pale, or sometimes showing areas of pale greyish red. Colonies on G25N 12–20 mm diam. At 58C, colonies of 1– 4 mm diam produced. At 378C, usually no growth, although in isolates from the tropics, colonies up to 35 diam Produced. On PDA, colonies of dense to floccose mycelium, white to pale salmon, becoming brown with age, with a central mass of orange to brown sporodochia, sometimes surrounded by poorly defined sporodochial rings; reverse pale salmon, often with a brown central area and brown flecks. On DCPA, colony appearance usually dominated by salmon, orange or brownish

Fig. 5.26 Fusarium equiseti (a) colonies on PDA and DCPA, 7 d, 258C; (b, c) macroconidia, bar ¼ 10 mm; (d) chlamydoconidia, bar ¼ 10 mm

102

sporodochia centrally and in poor to well defined concentric rings; mycelium low and sparse, coloured white or pale salmon; reverse pale. Macroconidia distinctly curved, often with a ‘‘hunchbacked’’ appearance, with five to seven septa, ranging from relatively short in some isolates to very long in others, with a wide range of sizes often present in a single isolate, basal cell distinctly foot-shaped, often with an elongated heel, apical cell elongated and curved, in long spores becoming filamentous; chlamydoconidia usually produced abundantly, in chains or clumps; microconidia absent. Distinctive features. Fusarium equiseti produces distinctly curved macroconidia which are often elongate, especially in the basal and apical cells. Colonies on PDA are floccose, usually with at least a central mass of orange sporodochia, and with reverse pale salmon, often flecked with brown. Chlamydoconidia are usually abundant; microconidia are not produced. Taxonomy. The teleomorph of Fusarium equiseti is Gibberella intricans Wollenw. However, there are few records of its occurrence in nature (Booth, 1971). Physiology. Fusarium equiseti grows strongly at 308C (Burgess et al., 1994) but, from observations in our laboratory, most isolates grow poorly or not at all at 378C. The minimum aw for growth has been reported to be 0.92 aw (Chen, 1966). Growth occurred at pH 3.3 but not 2.4, and was still rapid at pH 10.4 (Wheeler et al., 1991). Tripathi et al. (1999) reported growth between pH 2.0 and 11.00. Growth in N2 with <1% O2 was 90% of that in air (Hocking, 1990). Mycotoxins. Fusarium equiseti has been reliably reported to produce a number of mycotoxins. Trichothecene production in F. equiseti is variable, with both type A trichothecenes (diacetoxyscirpenol and related compounds, T-2 toxin, HT-2 toxin and neosolaniol) and type B trichothecenes (deoxynivalenol, 15-acetyldeoxynivalenol, fusarenone-X and nivalenol) reliably reported (see Pitt and Hocking, 1997; Langseth et al., 1999; Hestbjerg et al., 2002; Kosiak et al., 2005; Desjardins, 2006). Zearalenone, moniliformin, beauvericin, fusarochromanone and related compounds have also been reported (see Pitt and Hocking, 1997; Logrieco et al., 1998a; Desjardins, 2006), but production of butenolide is questionable (Desjardins, 2006).

5 Primary Keys and Miscellaneous Fungi

Ecology. A cosmopolitan soil fungus, Fusarium equiseti, has a distribution extending from Alaska to the tropics (Domsch et al., 1980). It has been isolated from a variety of plants, particularly cereals, where it may cause stem and root rots (Leslie and Summerell, 2006). F. equiseti has been identified as a minor component of maize ear rot in Europe, and head blight of wheat and associated grains in Europe and North American (Desjardins, 2006). It has been reported from various cereal grains including wheat, maize, barley, rye and rice (see Pitt and Hocking, 1997; Pitt et al., 1998a) and also from peanuts, walnuts, soybeans, cowpeas, sorghum, oilseeds and herbs (see Pitt and Hocking, 1997). It has been isolated from tomatoes (Okoli and Erinle, 1989) and capsicums (Adisa, 1983; Hashmi and Ghaffar, 1991), and has been reported as contributing to soft rot of bell peppers in India (Shukla and Sharma, 2000) and crown rot of bananas (Wallbridge, 1981; Jime´nez et al., 1993). It has also been implicated as a cause of rots of cucurbit fruits in contact with soil (Burgess et al., 1994), dry rots of potato tubers (El-Hassan et al., 2004) and soft rot of pumpkins (Elmer, 1996). F. equiseti has caused spoilage of UHT processed fruit juices, due to its ability to grow in very low O2 tensions (Hocking, 1990). References. Domsch et al. (1980) as Gibberella intricans; Nelson et al. (1983); Desjardins (2006); Leslie and Summerell (2006).

Fusarium graminearum Schwabe

Fig. 5.27

Teleomorph: Gibberella zeae (Schwabe) Petch Colonies on CYA filling the whole Petri dish, often to the lid, of dense, floccose mycelium coloured greyish rose, greyish yellow or paler; reverse usually orange red to greyish ruby, though sometimes pale brownish pink. Colonies on MEA filling the whole Petri dish, often reaching the lid at the margins at least, of dense to openly floccose mycelium, in shades of greyish rose and greyish yellow to golden brown; reverse orange brown to yellowish brown, sometimes paler at the margins. Colonies on G25N 20–30 mm diam, occasionally more. At 58C, colonies of 5–12 mm diam produced. No growth at 378C. On PDA, colonies filling the whole Petri dish, of dense, floccose mycelium coloured olive brown,

5.17 Genus Fusarium

103

Fig. 5.27 Fusarium graminearum (a) colonies on PDA and DCPA, 7 d, 258C; (b) Gibberella zeae perithecium and ascospores, bar ¼ 25 mm; (c) macroconidia, bar ¼ 10 mm

yellowish brown, reddish brown or pale salmon, or in combinations of those colours; sometimes with a central mass of red brown to orange sporodochia; reverse ruby to dark ruby centrally, sometimes violet brown. On DCPA, colony appearance dominated by salmon to orange sporodochia in concentric rings, overlaid by sparse, floccose, pale salmon mycelium. Macroconidia usually with five septa, sometimes less, thick walled, straight to moderately curved, with the ventral surface almost straight and a smoothly arched dorsal surface; basal cell distinctly foot-shaped, apical cell tapered; chlamydoconidia formed tardily in some isolates; microconidia absent. Distinctive features. On PDA, colonies of Fusarium graminearum are usually highly coloured, with

dense to floccose greyish rose to golden brown mycelium and dark ruby reverse. Macroconidia are relatively straight and thick-walled, with a foot-shaped basal cell. Microconidia are not produced. Taxonomy. The teleomorph of Fusarium graminearum is Gibberella zeae (Schwabe) Petch. Francis and Burgess (1977) described two populations of F. graminearum, which they termed Group 1 and Group 2. Group 2 isolates, the real F. graminearum, are responsible for head scab of wheat and also cause destructive ear rot of maize (Burgess et al., 1981; Desjardins, 2006; Leslie and Summerell, 2006). Group 1 isolates, which cause root and crown rot of wheat, were given species status and described as Fusarium pseudograminearum O’Donnell and T. Aoki (teleomorph Gibberella coronicola

104

T. Aoki and O’Donnell) by Aoki and O’Donnell (1999a, b). F. graminearum is homothallic and forms perithecia readily in nature and in culture on Carnation Leaf Agar (Leslie and Summerell, 2006), whereas F. pseudograminearum is heterothallic and does not. O’Donnell et al. (2004) proposed that the ‘‘F. graminearum clade’’ be split into nine different species, based on geographical and molecular data; however, the morphological distinctions between these species are small. This proposal has not been taken up by all Fusarium researchers (Leslie and Summerell, 2006). The complete genome of F. graminearum has been sequenced and is available (Broad Institute, 2003). Physiology. The optimal temperature for growth of Fusarium graminearum is between 24 and 268C on both liquid and solid media at pH 6.7–7.2 (Booth, 1971), and 258C on irradiated wheat grains (Ramirez et al., 2006a). The minimum aw for growth is close to 0.90 at 15–258C (Cuero et al., 1987; Ramirez et al., 2006a). The minimum pH for growth is temperaturedependent, near 3.0 at 25 and 378C, and 2.4 at 308C. The maximum is near 9.5 at 378C, but greater than 10.2 at the lower temperatures (Wheeler et al., 1991). Mycotoxins. Fusarium graminearum produces numerous mycotoxins: almost 50 toxic compounds have been reported. The most important economically are type B trichothecenes – deoxynivalenol (DON) and its derivatives 3-acetyldeoxynivalenol (3-AcDON) and 15-acetyldeoxynivalenol (15AcDON), nivalenol (NIV) and its derivatives and zearalenone (see Marasas et al., 1984; Pitt and Hocking, 1997; Desjardins, 2006; Leslie and Summerell, 2006). See Desjardins (2006) for a comprehensive review of trichothecene mycotoxins. Other toxins reported include aurofusarin, culmorins, fusarin C and steroids (see Pitt and Hocking, 1997; Leslie and Summerell, 2006). Type A trichothecenes, including T-2 toxin, HT-2 toxin, diacetoxyscirpenol and neosolaniol, have been reported from some isolates. However, production of these toxins by F. graminearum was queried by Nelson et al. (1983) and remains in doubt. Based on production of the different trichothecenes, Fusarium graminearum has been divided into two chemotypes, chemotype I for deoxynivalenol producers and chemotype II for nivalenol producers. Strains producing 3-acetyldeoxynivalenol are classified as chemotype IA and those producing 15-acetyldeoxynivalenol as chemotype IB (Ichinoe

5 Primary Keys and Miscellaneous Fungi

et al., 1983; Logrieco et al., 1988; Miller et al., 1991). North American strains are predominantly chemotype IB (Abbas et al., 1989b; Mirocha et al., 1989b; Miller et al., 1991; Abramson et al., 1993; Desjardins, 2006). This chemotype also predominates in the Ukraine (Leonov et al., 1990) and New Zealand (Lauren et al., 1992). Chemotype IA predominates in China (Miller et al., 1991; Zhang et al., 2007b), Argentina (Ramirez et al., 2006b) and Italy (Logrieco et al., 1988) while in Japan (Sugiura et al., 1990) and Poland (Visconti et al., 1990) both chemotypes IA and IB are commonly isolated. Chemotype II occurs only rarely in North America (Miller et al., 1991) but is more common elsewhere, including New Zealand (Lauren et al., 1992), Australia (Blaney and Dodman, 1988, 2002; Tan et al., 2004), Japan (Sugiura et al., 1990), Korea (Lee et al., 1986; Kim et al., 1993), Italy (Logrieco et al., 1988) and South Africa (Sydenham et al., 1991). Ramirez et al. (2006a) reported that deoxynivalenol production occurred most rapidly at 258C, but the maximum amount was produced at 308C. The aw range for deoxynivalenol production was 0.95– 0.995 aw. Llorens et al. (2004b) reported that 288C was optimal for deoxynivalenol production, but nivalenol and 3-acetyldeoxynivalenol were produced optimally at 20 and 158C, respectively. Zearalenone production was optimal at 208C (Llorens et al., 2004a). Other toxins produced by Fusarium graminearum include butenolide, culmorin, sambucinol, calonectrin and related compounds, fusarins and a number of other minor metabolites (Desjardins, 2006). Ecology. Fusarium graminearum is primarily a pathogen of gramineous plants, particularly wheat, in which it causes crown rot and head scab. Reports have come from Australia, Canada, the United States, Japan, Korea, South Africa and many European countries (see Pitt and Hocking, 1997; Desjardins, 2006; Leslie and Summerell, 2006). F. graminearum also causes cob rot and stalk rot of maize in many countries leading to DON and zearalenone formation in maize (see Pitt and Hocking, 1997; Leslie and Summerell, 2006). Occurrence of F. graminearum in barley has been less frequent, but it is believed to be one cause of gushing in beer (Niessen et al., 1992). Other sources include sugar beets (Bosch and Mirocha, 1992), soybeans (Clear et al., 1989; Jacobsen et al., 1995),

5.17 Genus Fusarium

sorghum (Onyike and Nelson, 1992) and triticale (Perkowski et al., 1988). F. graminearum has been reported to cause dry rot in potatoes (Ali et al., 2005) and postharvest rots in pumpkins (Elmer, 1996). Crown rot of bananas may also be due to F. graminearum (Wallbridge, 1981; Jime´nez et al., 1993). Quorn, a mycoprotein produced for human consumption in the United Kingdom, is produced from a strain of Fusarium which was originally identified as F. graminearum. This strain, which does not produce any trichothecene toxins, has since been assigned to a

105

different species, F. venenatum Nirenburg (O’Donnell et al., 1998). For reviews of this topic see Edwards (1993); Trinci (1994) and Wiebe (2004). References. Domsch et al. (1980) (as Gibberella zeae); Nelson et al. (1983); Desjardins (2006); Leslie and Summerell (2006).

Fusarium longipes Wollenw. & Reinking

Fig. 5.28

Colonies on CYA and MEA covering the whole Petri dish, plane, deep and floccose or sometimes

Fig. 5.28 Fusarium longipes (a) colonies on PDA and DCPA, 7 d, 258C; (b) macroconidia, bar ¼ 10 mm; (c) chlamydoconidia, bar ¼ 10 mm

106

funiculose, mycelium pale orange to pale red or dull reddish, reverse on CYA brownish orange to light brown, on MEA deep red to brownish red. On G25N, colonies 15–22 mm diam, of floccose, pale orange mycelium, reverse in similar colours. No germination at 58C. At 378C, colonies of variable size, up to 65 mm diam, of white to very pale pink mycelium, reverse dull yellow to pinkish brown. On PDA, colonies covering the whole Petri dish, similar to those on CYA, reverse dull red to dark red. On DCPA, colonies 55–65 mm diam, sparse, mycelium pale pink, showing abundant orange sporodochia containing macroconidia, reverse dull yellow to brown. Macroconidia on DCPA strongly curved, 50–80  4–6 mm, with four to six septa; apical cells usually with a long papilla or a long slender whip-like end; basal cells with a deep notch and long ‘‘foot’’. Microconidia not produced. Chlamydoconidia produced by some isolates, in chains or small clusters, spherical to ellipsoidal or irregular in shape, 7–12 mm wide, with thin smooth walls, becoming rough in age. Distinctive features. Pale orange to pale red mycelium on CYA and MEA, fast growth at 378C and very long, slender, strongly curved macroconidia are the main distinguishing features of Fusarium longipes. Taxonomy. No teleomorph is known for this species. Physiology. No studies on the physiology of this species are known to us. Mycotoxins. This appears to be a nontoxigenic species (Wing et al., 1993; Nelson et al., 1994), although Logrieco et al. (1998a) reported beauvericin production by one isolate. Ecology. Fusarium longipes principally comes from tropical soils (Burgess et al., 1994; Sangalang et al., 1995). It has been reported from peanuts from West Java (Dharmaputra and Retnowati, 1996), and we isolated it from 13% of Indonesian peanut samples, at up to 16% of kernels in infected samples and 1% of all kernels examined (Pitt et al., 1998a). It was also found at low levels in Indonesian maize and cowpeas, Thai maize, mung beans, cassava and sesame seed, and paddy rice from the Philippines (Pitt et al., 1993, 1998a). F. longipes has also been reported from cereals in Iran (Zare and Ershad, 1997). References. Nelson et al. (1983); Leslie and Summerell (2006).

5 Primary Keys and Miscellaneous Fungi

Fusarium oxysporum Schltdl.

Fig. 5.29

Colonies on CYA 50–70 mm diam, sometimes covering the whole Petri dish, moderately deep, of floccose white to greyish mycelium; reverse pale to pale greenish grey. Colonies on MEA 65–70 mm diam, often covering the whole Petri dish, of floccose white to pale greyish magenta mycelium; reverse greyish magenta to dark purple, often paler at the margins. Colonies on G25N 12–16 mm diam, occasionally larger. At 58C, germination to formation of microcolonies. At 378C, no growth, or colonies up to 5 mm diam formed. On PDA, colonies of white, pale salmon or pale mauve mycelium, sometimes dense and floccose, sometimes low, often with salmon sporodochia in a central mass, and sometimes in one or two poorly defined concentric rings also; reverse pale salmon, often mauvish centrally, sometimes dark magenta. On DCPA, colonies low, sometimes with concentric rings of sparse, floccose aerial mycelium, powdery with microconidia on the agar surface, or sometimes uniformly low with little aerial mycelium and then with appearance dominated by surface macroconidia. Macroconidia only slightly curved, usually with three septa, occasionally more, thin walled, with notched or foot shaped basal cells and short, sometimes hooked, apical cells; microconidia abundant, fusiform to kidney-shaped, produced in false heads from short, stout monophialides. Chlamydoconidia produced singly or in pairs. Distinctive features. Fusarium oxysporum produces abundant fusiform to kidney-shaped microconidia in false heads from short, stout, flaskshaped phialides in the aerial mycelium. The colony reverse on PDA is usually mauve, violet or greyish magenta. Taxonomy. Many plant pathogenic races exist in Fusarium oxysporum; these are called formae speciales (abbreviated f. sp.). Recent DNA sequence analyses have indicated that some of these are polyphyletic (Summerell et al., 2003). The most important of these are discussed by Booth (1971) and Leslie and Summerell (2006). No teleomorph is known. Physiology. Domsch et al. (1980) reported an optimum growth temperature between 25 and 308C for Fusarium oxysporum, with a minimum above 58C, and a maximum at or below 378C. The

5.17 Genus Fusarium

107

Fig. 5.29 Fusarium oxysporum (a) colonies on PDA and DCPA, 7 d, 258C; (b) phialides, bar ¼ 10 mm; (c) macroconidia and microconidia, bar ¼ 10 mm

optimum pH for growth is 7.7 with the wide range of pH 2.2–9.0 being tolerated (Domsch et al., 1980). The minimum aw for growth is 0.89 at 208C, after a germination time of 2 months (Schneider, 1954). Growth in N2 with <1% O2 was 97% of that in air (Hocking, 1990). Some growth occurred in 99% CO2, with only trace amounts of O2 and N2 present (Hocking, 1990). Atmospheres of 20–40% CO2 with 5 or 1% O2 reduced colony growth rate of F. oxysporum by 40–50%, but ergosterol content was reduced by 80–90% under these conditions (Taniwaki et al., 2001a). F. oxysporum chlamydoconidia do not exhibit exception heat resistance, despite its ability to cause spoilage in beverages (see below). In experiments in our laboratory, survival was recorded from mixed suspensions of microconidia and chlamydoconidia after heating at 638C for up to 48 s, but

not after exposure to 658C for 6 s in 208 Brix sucrose solution at pH 4.2. F. oxysporum was more resistant to dry heat, with sporadic survival detected after exposure to 1408C for 60 s (our unpublished data). Mycotoxins. Fusarium oxysporum has been widely reported to produce moniliformin (Marasas et al., 1979; Abbas et al., 1989b, 1991; Mirocha et al., 1989a; Chelkowski et al., 1990) and zearalenone (Per, 1995; see also Pitt and Hocking, 1997), although production of the latter compound appears to be in doubt as F. oxysporum does not possess the tri5 gene required for trichothecene biosynthesis (Tan and Niessen, 2003). F. oxysporum isolates reported to produce wortmannin (Abbas and Mirocha, 1988; Abbas et al., 1989a, 1992; Gunther et al., 1989) have been re-examined and classified as F. torulosum (Thrane, 2001). Other mycotoxins reported

108

to be produced by F. oxysporum are beauvericin (Logrieco et al., 1998a; Moretti et al., 2002), type B fumonisins (Abbas et al., 1995; Proctor et al., 2004) and type C fumonisins (Seo et al., 1996). Ecology. Fusarium oxysporum is a serious wilt pathogen of many crop plants, including sweet potatoes, cabbage and other crucifers, cucumbers and melons, oil and date palms, tomatoes, peas, soybeans and cowpeas, clovers, cotton and a variety of others (Booth, 1971; Nelson et al., 1981) and is geographically widespread (Sangalang et al., 1995; Leslie and Summerell, 2006). Considering these facts, its common occurrence in foods is not unexpected. It occurs in cereals, including maize in the United States, Australia, Turkey, South Africa and the Philippines; rice in India, Egypt and Indonesia; barley in Egypt and sorghum from Africa (see Pitt and Hocking, 1997). It is widespread on nuts: peanuts, pecans, hazelnuts and walnuts (see Pitt and Hocking, 1997). This species is one cause of crown rot of bananas (Wallbridge, 1981; Jime´nez et al., 1993; Boruah et al., 2004), and also occurs as a minor pathogen on citrus, pome fruits, tomatoes, melons (Snowdon, 1990) and pineapples (Biswal et al., 2007). Other sources include coriander seeds (Hashmi and Ghaffar, 1991), cacao beans (Ribeiro et al., 1986), navy beans (Tseng et al., 1996), adzuki and mung beans (Tseng and Tu, 1997), garlic (Rath and Mohanty, 1986), capsicums (Hashmi and Thrane, 1990), truffles (Bokhary et al., 1990), asparagus (Weber et al., 2006), cheese (Hocking and Faedo, 1992; Lund et al., 1995) and potatoes (Vrany et al., 1989; Kim and Lee, 1994; Venter and Steyn, 1998). F. oxysporum has been implicated as a cause of gill blackening disease in Kumma prawns (Penaeus japonicus) (Souheil et al., 1999). Due to its ability to grow in low O2 tensions, Fusarium oxysporum has caused spoilage of UHTprocessed fruit juices (Hocking, 1990). This problem became widespread from 2001 onwards, and our laboratory investigated several spoilage incidents occurring in Australia and New Zealand in fruit, dairy- and soy-based aseptically filled beverages. We had anecdotal evidence of similar problems in Europe, Asia and North America. F. oxysporum appeared to be colonising the aseptic area of the packaging equipment, and once established, was very difficult to eliminate.

5 Primary Keys and Miscellaneous Fungi

References. Booth (1971); Domsch et al. (1980); Nelson et al. (1981, 1983); Leslie and Summerell (2006).

Fusarium poae (Peck) Wollenw.

Fig. 5.30

Colonies on CYA filling the whole Petri dish, deep, of moderately dense white to pale rose mycelium; reverse unevenly coloured, pale to rose red or deep red. Colonies on MEA filling the whole Petri dish, deep, of sparse, pale rose mycelium; reverse brownish orange or paler. Colonies on G25N 8–12 mm diam. At 58C, germination to colonies up to 2 mm diam formed. No growth at 378C. On PDA, colonies moderately deep, of floccose, pale salmon to pale rose mycelium, darker centrally, reverse varying from pale salmon at the margins to greyish ruby centrally, or entirely dark ruby to dark magenta. On DCPA, colonies moderately deep, of floccose pale salmon mycelium in poorly defined concentric rings; sporodochia rarely present; reverse pale or, in highly pigmented isolates, with annular brownish red rings. Macroconidia usually sparsely produced, varying in shape, mostly with three septa, occasionally more, slightly curved, with a foot-shaped basal cell. Microconidia abundant, spherical, often with a distinct papilla, occasionally also lemon-shaped, aseptate or with one septum, produced in the aerial mycelium from short flask-shaped phialides on compact branched stipes, often appearing like bunches of grapes when examined in situ under the low power microscope. Distinctive features. The abundant production of spherical microconidia borne from flask-shaped phialides on compact, branched stipes distinguishes Fusarium poae from other species. It also has a distinctive odour, described as ‘‘fruity’’ or ‘‘sweet’’ (Leslie and Summerell, 2006). Taxonomy. A group of cultures morphologically similar to Fusarium poae, referred to as ‘‘powdery’’ F. poae, have recently been assigned to a separate species Fusarium langsethiae Torp and Nirenburg (Torp and Nirenburg, 2004). This species has been isolated from oats, wheat and barley in cooler areas of Europe and Scandinavia. F. langsethiae, which does not have the characteristic sweet odour of F. poae, is also distinguished by its metabolic profile (Thrane et al., 2004) and by

5.17 Genus Fusarium

109

Fig. 5.30 Fusarium poae (a) colonies on PDA and DCPA, 7 d, 258C; (b) macroconidia, bar ¼ 10 mm; (c) microconidia, bar ¼ 10 mm; (d) phialides, bar ¼ 10 mm

molecular techniques (Schmidt et al., 2004). It appears to be more closely related to F. sporotrichioides than F. poae (Niessen et al., 2004). Physiology. The optimum temperature for growth of Fusarium poae is near 258C, with a minimum near 2.58C and a maximum of 358C (Kvashnina, 1976; Torp and Nirenburg, 2004). The minimum aw for growth is near 0.90 between 17 and 258C (Magan and Lacey, 1984c). Mycotoxins. Reports of toxin production by Fusarium poae have been very variable. Reports

that it was perhaps responsible for Alimentary Toxic Aleukia due to the production of T-2 toxin have largely been discounted, as F. sporotrichioides, the major producer of that toxin, was also commonly present during that disease syndrome, and in the haemorrhagic diseases that occurred in cattle, pigs and poultry in the United States in the 1960s (Marasas et al., 1984). However, it has been concluded more recently that certain strains of F. poae can indeed produce T-2 toxin (De Nijs et al., 1996; Langseth et al., 1999; Thrane et al., 2004).

110

Most strains of Fusarium poae produce nivalenol, diacetoxyscirpenol, monoacetoxyscirpenols, fusarenone-X and scirpentriol (Thrane et al., 2004), with some strains also producing beauvericin and enniatins (Thrane et al., 2004; Chelkowski et al., 2007). Logrieco et al. (1998a) also reported beauvericin production by F. poae. T-2 toxin is more often produced by F. sporotrichioides and F. langsethiae (Thrane et al., 2004; Desjardins, 2006). Ecology. Fusarium poae mainly occurs in temperate regions, where it is found on woody seedlings or herbaceous and gramineous hosts (Booth, 1971; Leslie and Summerell, 2006). From foodstuffs, it has been reported most commonly from grains, especially wheat, from France, Canada, Japan, Ethiopia, Hungary, Poland, Germany and Norway (see Pitt and Hocking, 1997; Birzele et al., 2002; Bottalico and Perrone, 2002; Ioos et al., 2004; Kosiak et al., 2003, 2004). It has been implicated as a cause of panicle blight of oats in Poland (Mielniczuk, 2001), head blight of oats in Canada (Tekauz et al., 2004) and as a pathogen of barley in Argentina (Barreto et al., 2004). It has been associated with maize in cooler climates, where it may be involved in ear rot (see Pitt and Hocking, 1997; Logrieco et al., 2002; Chelkowski, et al., 2007). F. poae has also been recorded from soybeans in the United States (Abbas and Bosch, 1990), heart rot of sugar cane in South Africa, rice in Australia and decayed stored citrus fruit in Georgia, CIS (Booth, 1971). References. Booth (1971); Domsch et al. (1980); Nelson et al. (1983); Desjardins (2006); Leslie and Summerell (2006).

Fusarium proliferatum (Matsush.) Nirenberg Fig. 5.31 Teleomorph: Gibberella intermedia (Kuhlman) Samuels Colonies on CYA covering the whole Petri dish, plane, floccose, of pale orange mycelium, reverse light orange to pastel red. Colonies on MEA similar to those on CYA but much less dense, mycelium in similar colours, reverse orange, greyish red or violet brown. On G25N, germination or colonies up to 5 mm diam. At 58C, no germination. At 378C, colonies 5–20 mm diam, of pale orange mycelium, reverse usually orange.

5 Primary Keys and Miscellaneous Fungi

On PDA, colonies covering the whole Petri dish, similar to on MEA, reverse light orange to greyish red. On DCPA, colonies 45–55 mm diam, of pale orange mycelium, plane, floccose, similar in texture to on CYA, orange sporodochia usually evident on the agar surface, reverse orange brown. Macroconidia on DCPA slightly curved, with three to four septa, 40–50  3.0–4.0 mm, rounded or papillate at both ends, or sometimes with a notch at the basal end, with thin, smooth walls. Microconidia borne in chains and false heads from monophialides or polyphialides, ellipsoidal or beanshaped, commonly 8–12  2.5–4.0 mm and nonseptate, occasionally longer and with one or more septa, with thin smooth walls. Chlamydoconidia not produced. Distinctive features. Similar to Fusarium verticillioides in most respects, F. proliferatum is distinguished by the formation of microconidia from polyphialides. Microconidial chains are usually shorter in F. proliferaturm than F. verticillioides. Freshly isolated cultures form abundant macroconidia in sporodochia, but this feature is often lost after repeated transfer (Leslie and Summerell, 2006). Taxonomy. Morphologically, Fusarium proliferatum is indistinguishable from F. fujikuroi Nirenburg. These two species can be differentiated using mating tests or by DNA sequencing. F. fujikuroi, which is associated mainly with rice, does not produce fumonisins, whereas most isolates of F. proliferatum do. Physiology. The optimum pH and temperature for growth of Fusarium proliferatum are 5.5 and 258C, respectively, with growth occurring at 5 and 378C, but not 408C on maize meal (Marı´ n et al., 1996). The minimum aw for germination reported by Marı´ n et al. (1996) was 0.88, with no germination occurring at 0.85 aw after 40 days. However, Samapundo et al. (2005a) reported a minimum aw of 0.87 for growth of F. proliferatum on maize, with optimum growth occurring at 308C at 0.97 aw, the highest aw tested. F. proliferatum was able to grow on irradiated maize kernels at 0.98 and 0.93 aw in atmospheres containing up to 60% CO2 with 20% O2, although the lag time was longer and the growth rate slower than the controls (Samapundo et al., 2007a). Reduction of headspace oxygen from 20 to 2% (without elevated CO2) had no effect on growth of F. proliferatum, but

5.17 Genus Fusarium

111

Fig. 5.31 Fusarium proliferatum (a) colonies on PDA and DCPA, 7 d, 258C; (b) phialides bearing microconidia in chains and false heads in situ, bar ¼ 50 mm; (c) macroconidia and microconidia, bar ¼ 10 mm; (d) polyphialides, bar ¼ 10 mm; (e) monophialides, bar ¼ 10 mm

vacuum packaging and the incorporation of oxygen scavenging sachets prevented growth (Samapundo et al., 2007b). Application of 1% ammonium bicarbonate to maize at aw values between 0.99 and 0.92 prevents growth and fumonisin production by F. proliferatum and was proposed as a possible treatment for stored maize (Samapundo et al., 2007c). The antioxidants propylparaben and butylated

hydroxyanisole (BHA) can also inhibit growth and fumonisin production by F. proliferatum under some conditions (Etcheverry et al., 2002). Fumonisin B1 production was inhibited at headspace CO2 concentrations of 40, 30 and 10% at 0.98, 0.95 and 0.93, respectively (Samapundo et al., 2007a). When headspace oxygen was controlled, optimum fumonisin production occurred

112

with 20% O2 at 0.98 aw, but at 0.95 and 0.93 aw, more fumonisin was produced with 10% headspace O2 (Samapundo et al., 2007b). Marı´ n et al. (1995) reported fumonisin production was higher at 25 than at 308C, and highest at the highest aw tested (0.97), with low levels still produced at 0.92 aw. Samapundo et al. (2005b) found that there was little difference in fumonisin production at 25 and 308C, but lower amounts were produced at 158C, irrespective of aw value (between 0.97 and 0.92). Conversely, Melcion et al. (1998) observed maximal fumonisin production at 158C on maize at very high aw. Mycotoxins. Fusarium proliferatum is a major producer of fumonisins B1, B2 and B3 (Ross et al., 1990; see also Pitt and Hocking, 1997; Desjardins, 2006), and indeed strains of F. proliferatum are among the highest fumonisin producers (Leslie et al., 2004). Other mycotoxins produced include moniliformin, beauvericin, enniatins and fusarin (Marasas et al., 1986; Desjardins et al., 2000a; see also Desjardins, 2006). Ecology. Reporting of Fusarium proliferatum in foods and feeds has increased sharply with the discovery of fumonisins and that this species is an important source of these toxins. It has been shown to colonise maize plants throughout the world, and is increasingly important in maize ear rot in Europe (see Bacon and Nelson, 1994; Desjardins, 2006; Leslie and Summerell, 2006). It was reported from sorghum in Nigeria by Onyike and Nelson (1992) and in pine nuts in Spain (Marı´ n et al., 2007b). We have encountered this species in maize, mung beans, paddy rice and occasionally sorghum in Southeast Asia (Pitt et al., 1998a). F. proliferatum has also been reported as a cause of black point in wheat (Conner et al., 1996; Desjardins et al., 2007). It is a pathogen of bananas (Jime´nez et al., 1993), onions (Toit et al., 2003; Stankovic et al., 2007), garlic (Dugan et al., 2003, 2007; Stankovic et al., 2007) and asparagus (Elmer, 2000; Gossman et al., 2005; Corpas-Hervias et al., 2006; Weber et al., 2006; Liu et al., 2007). Fumonisins have been reported in infected garlic (Seefelder et al., 2002) and asparagus (Logrieco et al., 1998b; Seefelder et al., 2002; Weber et al., 2006; Liu et al., 2007). References. Nelson et al. (1983); Desjardins (2006); Leslie and Summerell (2006).

5 Primary Keys and Miscellaneous Fungi

Fusarium semitectum Berk. & Ravenel Fig. 5.32 Fusarium incarnatum (Desm.) Sacc. Fusarium pallidoroseum (Cooke) Sacc.

Colonies on CYA filling the whole Petri dish, sometimes to the lid at the edges, of dense, floccose, white, pale salmon or pale brown mycelium, sometimes powdery from macroconidia produced in the aerial mycelium; pale salmon sporodochia occasionally produced at the inoculation points; reverse pale salmon, yellowish, greyish yellow or with brown patches. Colonies on MEA filling the whole Petri dish, of floccose or funiculose white to pale greyish yellow mycelium; reverse pale, sometimes greyish orange centrally or in patches. Colonies on G25N 15–20 mm diam. At 58C, usually only microcolonies produced. Usually no growth at 378C, but occasionally colonies of 2 mm diam formed. On PDA, colonies of dense, floccose mycelium, coloured pale salmon, brown or yellow, sometimes with a powdery appearance from macroconidia in the aerial mycelium; reverse pale salmon, often brownish centrally. On DCPA, colonies in similar colours to those on PDA, but less dense, aerial mycelium often powdery with macroconidia, and occasionally a central area of salmon to orange sporodochia; reverse pale. Macroconidia of two types produced: the first in the aerial mycelium, often from polyphialides, cigaror spindle-shaped, straight or slightly curved, with four to five septa, and with poorly differentiated basal and apical cells; the second in sporodochia, more curved, with a foot-shaped basal cell and curved apical cell. Chlamydoconidia usually produced; microconidia not produced. Distinctive features. The production of spindleor cigar-shaped macroconidia from polyphialides in the aerial mycelium is the most distinctive feature of Fusarium semitectum. These macroconidia can be observed in situ with the low power microscope, and are often arranged in pairs in ‘‘V’’ shapes, resembling rabbit ears. Taxonomy. Type material of Fusarium semitectum, examined by Booth and Sutton (1984), was found to be a Colletotrichum species, C. musae. However, type material of F. pallidoroseum (Cooke) Sacc., which they also examined, contained polyphialides and macroconidia typical of F. semitectum. Subsequently, Nirenberg (1990) pointed out that the species epithet incarnatum predates pallidoroseum by about 30

5.17 Genus Fusarium

113

Fig. 5.32 Fusarium semitectum (a) colonies on PDA and DCPA, 7 d, 258C; (b) phialides bearing macroconidia in aerial mycelium in situ, bar ¼ 50 mm; (c) phialides, bar ¼ 10 mm; (d) macroconidia, bar ¼ 10 mm

years, so if the name were to be changed, it should be F. incarnatum. However, both Desjardins (2006) and Leslie and Summerell (2006) agree that F. semitectum is really a species complex, with possibly five intra-specific groups (Abd-Elsalam et al., 2003), and until such time as it is thoroughly examined both morphologically and phylogenetically, the name F. semitectum should continue to be used. Physiology. The temperature range for growth of Fusarium semitectum is 38C to ca. 378C, with the optimum near 258C (Kakker and Mehrotra, 1971;

Marin et al., 1996; Shukla and Sharma, 2000). Cherian (2007) reported maximum rotting of bananas by F. semitectum occurred at 308C. Mycotoxins. Confusion still exists over the mycotoxins produced by Fusarium semitectum (Desjardins, 2006). Zearalenone production has been reported (see Marasas et al., 1984; Pitt and Hocking, 1997, Desjardins, 2006, Leslie and Summerell, 2006). Production of T-2 toxin has also been reported, but is unsubstantiated (Marasas et al., 1984; Desjardins, 2006). Japanese isolates of F.

114

semitectum have been reported to produce diacetoxyscirpenol, nivalenol, fusarenone-X and neosolaniol (Suzuki et al., 1980, 1981). Diacetoxyscirpenol production has also been reported by Molto et al. (1997). Ecology. Fusarium semitectum is widespread in tropical and subtropical countries, although reports from foods are relatively uncommon. It has been found at low levels in maize in Italy (Logrieco et al., 1995) and the USA (Katta et al., 1995), and we have found high levels of F. semitectum in Thai maize (45% of samples, up to 22% infected grains in infected samples and 4% of all kernels examined; Pitt et al., 1993). Incidence in Indonesian maize was similar, but levels in Philippine maize lower (Pitt et al., 1998a). F. semitectum is one of the dominant fungi of pearl millet (Wilson, 2002; Jurjevic et al., 2007) and is common in peanuts (Joffe, 1969; Gilman, 1969; Oyeniran, 1980; Pitt et al., 1993; Dharmaputra and Retnowati, 1996). It has also been reported from rice (Desjardins et al., 2000a) and sorghum (Onyike and Nelson, 1992; Usha et al., 1994; Shabbir and Rajasab, 2004). It causes storage rots of various fruits (Snowdon, 1990), especially crown rot of bananas (see Pitt and Hocking, 1997; Vesonder et al., 1995; Marin et al., 1996; Cherian, 2007) and soft rot of bell peppers (Shukla and Sharma, 2000; Sharma and Shukla, 2003). It has also been reported to cause postharvest spoilage of tomatoes (Muniz et al. 2003; Chaturbhuj and Rai, 2005), passionfruit (Muniz et al., 2003), mushrooms (Kang et al., 2002) and disease in walnuts (Belisario et al., 1999). This species has also been isolated from pigeon peas (Maximay et al., 1992), soybeans (Vaamonde et al., 1987; and our unpublished data), black gram (Goyal and Jain, 1998), black beans (Castillo et al., 2004), coriander (Hashmi and Thrane, 1990; Hashmi and Ghaffar, 1991) and sunflower seeds (Shahnaz and Ghaffar, 1991), though with few reports in each case. Earlier reviews indicate isolations from citrus fruits, tomatoes, melons, cucumbers and potatoes (Booth, 1971; Domsch et al., 1980). In our experience, Fusarium semitectum is much more common in tropical commodities than the above citations would indicate. This species appears to be endemic in paddy rice from Thailand (in 94% of samples examined, up to 44% of grains infected in infected samples and a very high 20% of all grains

5 Primary Keys and Miscellaneous Fungi

examined; Pitt et al., 1994) and Indonesia (60% of samples, up to 60% of grains in infected samples and 7% of all grains; Pitt et al., 1998a). It was the most common fungus in Thai mung beans (55% of samples; up to 76% infection in infected samples, and 15% infection overall). It was also common in Thai black beans (6% of all beans examined), sorghum (4% of all grains) and soybeans (2% of all beans; Pitt et al., 1994). Generally similar figures were found for infection in rice, soybeans, black beans and mung beans in Indonesia and the Philippines (Pitt et al., 1998a). Indeed, F. semitectum was by far the most common fungal invader of beans that we encountered in Southeast Asia. References. Domsch et al. (1980); Nelson et al. (1983); Desjardins (2006); Leslie and Summerell (2006).

Fusarium solani (Mart.) Sacc.

Fig. 5.33

Teleomorph: Haemanectria haematococca (Berk. & Broome) Samuels & Nirenberg Colonies on CYA 60–65 mm diam, low, of moderately dense white mycelium, often covered in very fine droplets of clear exudate; a central, cream spore mass sometimes present; reverse pale, sometimes with bluish or greenish areas. Colonies on MEA 50–60 mm diam, low to moderately deep, of sparse, often slightly funiculose, white to pale violet mycelium; reverse pale, sometimes bluish grey centrally. Colonies on G25N 3–8 mm diam. No growth at 58C. At 378C, either no growth, or colonies up to 10 mm diam formed. On PDA, colonies low to moderately deep, of white to cream mycelium in concentric rings, often alternating with rings of cream or bluish grey sporodochia; in some isolates the teleomorph produced, of dark orange perithecia scattered over the central area of the colony; reverse pale or with areas of turquoise grey or pale violet brown. On DCPA, colonies low, of sparse, white mycelium in annular rings, with a central mass of cream sporodochia; reverse pale to pale yellow brown. Macroconidia abundant, stout, thick walled, with three to four, or less commonly five septa, straight, parallel sided for most of the length, apical cells blunt and rounded, basal cells either rounded, notched or sometimes distinctly foot-shaped.

5.17 Genus Fusarium

115

Fig. 5.33 Fusarium solani (a) colonies on PDA and DCPA, 7 d, 258C; (b) phialides bearing macroconidia in false heads in situ, bar ¼ 50 mm; (c) phialides, bar ¼ 10 mm; (d) microconidia, bar ¼ 10 mm; (e) macroconidia, bar ¼ 10 mm

Microconidia usually abundant, ellipsoidal, fusiform or kidney-shaped, produced in false heads on very long, straight phialides. Chlamydoconidia produced singly or in pairs. Distinctive features. Fusarium solani is the only Fusarium species which produces cream or bluish rather than salmon or orange sporodochia. The macroconidia are stout and straight or only slightly curved. Microconidia are produced in false heads (mucoid balls) on very long, slender phialides. This species is morphologically very similar to

F. oxysporum. However, phialides in F. solani are much longer than those of F. oxysporum. Taxonomy. The teleomorph of Fusarium solani is Haemanectria haematococca (Berk. and Broome) Samuels and Nirenberg, which produces orange brown perithecia. However, only about 1% of F. solani isolates produce the teleomorph in culture (Booth, 1983). Leslie and Summerell (2006) believe that there are a number of biological species within F. solani and that further work is required to clarify the taxonomy and phylogeny of this group.

116

Physiology. Domsch et al. (1980) noted that various authors have reported the optimum growth temperature for Fusarium solani to be between 27 and 318C, with strong growth at 378C. However, our observations indicate only weak growth at this temperature, and Chaturvedi et al. (2003) reported optimum growth at 258C. Schneider (1954) reported growth of F. solani down to 0.90 aw after a germination time of 8 weeks at 208C. Mycotoxins. Mouldy sweet potatoes (Ipomoea batatas) or extracts from them have been shown to be toxic to a variety of animals, including cattle (Wilson et al., 1970; Doupnik et al., 1971), chickens (Doupnik et al., 1971; Peckham et al., 1972) and mice (Boyd and Wilson, 1972). The toxic principles are believed to be furanoterpenoids, ipomeanols and ipomeanine (Nelson et al., 1983, 1994; Mawhinney et al., 2008). These compounds are not mycotoxins, but result from fungal metabolism of phytoalexins produced by the plant (see Leslie and Summerell, 2006). Fusarium solani has been reported to produce fusaric acid (Bacon et al., 1996) and the immunosuppressive compound cyclosporin (Sugiura et al., 1999). Ecology. A cosmopolitan soil fungus, Fusarium solani, has frequently been isolated from subterranean crops such as potatoes, sweet potatoes, yams and peanuts (see Pitt and Hocking, 1997; Ismail, 2001; Peters et al., 2008). Hot water dipping (57.58C, 20–30 min) can reduce the incidence of storage rots of potatoes caused by F. solani (Ranganna et al., 1998). F. solani may also invade a wide variety of other vegetable crops (Snowdon, 1991), especially cucurbits including muskmelons (Champaco et al., 1993) and squash (Assawah and AlZarari, 1984), and legumes such as beans, soybeans and peas (see Pitt and Hocking, 1997; Tseng and Tu, 1997). Other sources are diverse, including small grains, where it may be part of the cohort of Fusarium species causing head blight (Bottalico and Perrone, 2002), maize, sorghum, bananas, guavas, cassava, sugar beets, capsicums, garlic, and seeds of sunflower, sesame and coriander (see Pitt and Hocking, 1997). We have isolated F. solani from beer, shortening flakes, chilled water lines in a beverage production plant, the inside of a juice filling machine, and an air conditioning duct. This species was widespread in Indonesian crops, especially beans, usually at about 1% of all seeds examined, in paddy rice, maize, peanuts and pepper (Pitt et al., 1998a).

5 Primary Keys and Miscellaneous Fungi

Fusarium solani can attack keratin, and has been isolated from human fingernails and eyes (Domsch et al., 1980; de Hoog et al., 2000; Leslie and Summerell, 2006). It can be a serious pathogen of crustaceans, attacking and destroying the chitinous exoskeleton (Fisher et al., 1978; Gonzalez, 1995; Le et al., 2005) and has also been reported to cause disease in sea turtles (Caban˜es et al., 1997; Castella´ et al., 1999a; de Hoog et al., 2000). References. Domsch et al. (1980); Nelson et al. (1983); Desjardins (2006); Leslie and Summerell (2006).

Fusarium sporotrichioides Sherb.

Fig. 5.34

Colonies on CYA filling the whole Petri dish, deep, of floccose white to pale pink and brown mycelium; reverse pale salmon centrally, brownish at the margins. Colonies on MEA similar to those on CYA except more highly coloured; reverse violet brown centrally, paler at the margins. Colonies on G25N 12–15 mm diam. At 58C, colonies 5–10 mm diam. No growth at 378C. On PDA, colonies of dense, floccose, salmon to pink mycelium, sometimes with a central mass of orange sporodochia; reverse greyish rose to burgundy, occasionally paler. On DCPA, colonies of floccose pale salmon mycelium in concentric rings; orange sporodochia sometimes present on the agar surface; reverse pale. Macroconidia abundant, moderately curved, with three to five septa and with a curved, pointed apical cell and a notched or foot-shaped basal cell. Microconidia abundant, produced from polyphialides in the aerial mycelium, fusiform, broadly ellipsoidal or pyriform, the latter often produced only on PDA, often with a papilla at the base. Chlamydoconidia formed abundantly, singly or in chains or clumps, as cultures age. Distinctive features. On PDA, Fusarium sporotrichioides has a similar appearance to F. poae, F. chlamydosporum and F. tricinctum. The production of both fusiform and pyriform microconidia from polyphialides distinguishes F. sporotrichioides from these closely related species. Pyriform microconidia are produced more commonly, and sometimes exclusively, on cultures on PDA. Colony reverses on PDA are always greyish rose or burgundy. Physiology. Domsch et al. (1980) gave the optimum growth temperature of Fusarium sporotrichioides as 22.5–27.58C, with a maximum of 358C. Joffe (1962)

5.17 Genus Fusarium

117

Fig. 5.34 Fusarium sporotrichioides (a) colonies on PDA and DCPA, 7 d, 258C; (b, c) polyphialides, bar ¼ 10 mm; (d) microconidia, bar ¼ 10 mm; (e) macroconidia, bar ¼ 10 mm

reported the growth of toxigenic isolates of this species down to –28C. Schneider (1954) reported 0.88 aw as the minimum for growth, after 8 weeks incubation at 208C. Mycotoxins. Fusarium sporotrichioides is regarded as the major cause of alimentary toxic aleukia (ATA) a devastating disease which occurred in the USSR during and after World War II, in times of extreme food shortage resulting in consumption of overwintered cereals (Joffe, 1978). Hundreds of thousands of people died from T-2 toxin produced by this species and F. poae (Marasas et al., 1984). F. sporotrichioides also produces a number of T-2 toxin derivatives, including HT-2, T-2 triol and T-2 tetraol, and biosynthetic intermediates such as neosolaniol, diacetoxyscirpenol and 15-monoacetoxyscirpenol (Thrane et al., 2004;

Abramson et al., 2004; see also Pitt and Hocking, 1997; Desjardins, 2006; Leslie and Summerell, 2006). F. sporotrichioides has been used to elucidate the biochemistry and genetics of trichothecene biosynthesis (Desjardins et al., 1993), with the first trichothecene biosynthetic gene, trichodiene synthase (TRI5) cloned from F. sporotrichioides in 1989 (see Desjardins, 2006; Leslie and Summerell, 2006). Early reports of deoxynivalenol and nivalenol production have not been confirmed and may be due to misidentification of isolates (Desjardins, 2006). Other mycotoxins produced by F. sporotrichioides include aurofusarin, beauvericin and occasionally enniatins, but not nivalenol, fusarenone-X or fumonisins (Thrane et al., 2004; Desjardins, 2006).

118

Fusarium sporotrichioides has been implicated as the cause of mouldy corn toxicosis (haemorrhagic syndrome) in cattle, pigs and poultry in the USA and elsewhere, fescue foot in cattle feeding on winter pastures in the USA, Australia and New Zealand, akakabi-byo (scabby grain intoxication) and bean hulls poisoning in animals in Japan (Marasas et al., 1984; Wu et al., 1997). Ramakrishna et al. (1996) reported that T-2 toxin formation by F. sporotrichioides on barley grain occurred optimally at 208C and toxin production was stimulated by the presence of Aspergillus flavus or Penicillium verrucosum. Ecology. While not so commonly isolated as some other species described here, Fusarium sporotrichioides is important because of its toxicity. It occurs more commonly in cool climates and, in foods, is almost entirely confined to grains. Historically, this species has been associated with rye in Europe (Joffe, 1978; Matossian, 1989). More recently, it has been reported most frequently from wheat, in Japan, Spain, Hungary, Poland, Canada, Ethiopia (see Pitt and Hocking, 1997) and Finland and Russia (Yli-Mattila et al., 2004). Other grain sources include maize (Logrieco et al., 1995, 2002; Adejumo et al., 2007; and see Pitt and Hocking, 1997), barley (Yli-Mattila et al., 2004; Bourdages et al., 2006), sorghum, wild rice (Nyvall et al., 1999) and cereal grains in general (see Pitt and Hocking, 1997; Langseth et al., 1998; Kosiak et al., 2003). This species has also been isolated from peanuts (Austwick and Ayerst, 1963), sugar beets (Bosch and Mirocha, 1992) and in our laboratory from Australian soybeans. References. Domsch et al. (1980); Nelson et al. (1983); Desjardins (2006); Leslie and Summerell (2006). Fusarium subglutinans (Wollenw. & Reinking) P.E. Nelson et al. Fig. 5.35 Fusarium moniliforme J. Sheld. var. subglutinans Wollenw. & Reinking Fusarium sacchari (Butler) W. Gams var. subglutinans Wollenw. & Reinking

Teleomorph: Gibberella subglutinans (Edwards) P.E. Nelson et al. Colonies on CYA covering the whole Petri dish, of dense white to very pale salmon mycelium, in degenerate cultures of low and sparse funiculose white mycelium; sometimes with a central orange

5 Primary Keys and Miscellaneous Fungi

spore mass; reverse pale, salmon or yellowish. Colonies on MEA less dense than on CYA, of white to pale salmon mycelium, sometimes powdery with microconidia; reverse pale yellow, salmon or violet grey. Colonies on G25N 5–12 mm diam, occasionally larger. At 58C, germination to microcolony formation. No growth at 378C. On PDA, colonies low to moderately deep, of floccose to funiculose mycelium coloured white, pale salmon, pale pink or mauve, sometimes powdery with microconidia, reverse violet grey to deep violet centrally, paler at the margins, or uniformly pale. On DCPA, colonies low at the margins, floccose centrally, of pale salmon mycelium, sometimes overlying a thin layer of pale orange macroconidia on the agar surface; reverse pale. Macroconidia slightly curved to almost straight, with three to five septa and thin, delicate walls, narrow, tapered apical cells and foot-shaped basal cells. Microconidia abundant, fusiform or broadly ellipsoidal, aseptate or with a single septum, produced in false heads from polyphialides and also from simple phialides. Chlamydoconidia not produced. Distinctive features. The production of microconidia in false heads from polyphialides and the absence of chlamydoconidia distinguish Fusarium subglutinans from otherwise similar species such as F. verticillioides and F. oxysporum. Taxonomy. Fusarium subglutinans was raised to species status by Nelson et al. (1983), previously having been regarded as a variety of F. moniliforme or F. sacchari. The teleomorph of F. subglutinans is Gibberella subglutinans (Edwards) P.E. Nelson et al. Recent phylogenetic studies indicate that there may be at least two cryptic species within F. subglutinans (Steenkamp et al., 2002; Desjardins et al., 2006). Physiology. Martelleto et al. (1998) reported growth of Fusarium subglutinans between 10 and 308C, with 258C being optimal. Castella´ et al. (1999b) found growth of F. subglutinans was optimal between 20 and 258C on maize cultures, but on rice, growth was best at 158C. Highest production of fusaproliferin was at 208C on maize after 6 weeks (Castella´ et al., 1999b). Mycotoxins. Usually regarded as one of the less toxic of major Fusarium species, F. subglutinans has, however, been reported to produce moniliformin as its major toxin (see Pitt and Hocking, 1997; Kostecki et al., 1999; Sewram et al., 1999; Logrieco

5.17 Genus Fusarium

119

Fig. 5.35 Fusarium subglutinans (a) colonies on PDA and DCPA, 7 d, 258C; (b) polyphialides, bar ¼ 10 mm; (c) macroconidia amd microconidia, bar ¼ 10 mm

et al., 2002; Desjardins, 2006; Desjardins et al., 2006; Leslie and Summerell, 2006; Sørensen et al., 2007). Production of fumonisin B1 has also been reported (Visconti and Doko, 1994), but levels are generally low (Reynoso et al., 2004). Other toxins reported are beauvericin and fusaproliferin (Moretti et al., 1995; Logrieco et al., 1998a; Castella´ et al., 1999b; Kostecki et al., 1999; Shephard et al., 1999; Torres et al., 2001; Reynoso et al., 2004). Ecology. Closely related to Fusarium verticillioides, F. subglutinans appears to have a similar host range, and a worldwide distribution. Its major food habitat is maize. It has been isolated from Europe (Italy, Logrieco et al., 1995; Austria, Lew et al., 1991; Poland, Logrieco et al., 1993b), the United States (Abbas et al., 1988, 1989b; Katta

et al., 1995), Canada (Neish et al., 1983), the Caribbean (Julian et al., 1995), Peru (Logrieco et al., 1993a), Argentina (Torres et al., 2001; Reynoso et al., 2004, 2006), Mexico (Desjardins et al., 2000b; Morales-Rodriguez et al., 2007), South Africa (Marasas et al., 1978, 1979; Rheeder et al., 1995), Zimbabwe (Mubatanhema et al., 1999), Indonesia (Pitt et al., 1998a), Australia (Burgess et al., 1981; Blaney et al., 1986) and New Zealand (Hussein et al., 1991, 2002). F. subglutinans has also been isolated in Mexico from teosintes, the nearest wild relatives of maize (Desjardins et al., 2000b). F. subglutinans has been reported from wheat in Europe (Bottalico and Perrone, 2002; Ioos et al., 2004; Cosic et al., 2007), South Africa (Boshoff et al., 1998), Tanzania (van Dyk, 2004), Egypt (Gherbawy et al., 2006) and Iran (Moosawi-Jorf

120

et al., 2007), but it does not appear to play an active role in head blight. Fusarium subglutinans is a pathogen of pineapples (Bolkan et al., 1979; Rohrbach and Taniguchi, 1984, Matos et al., 2000), bananas (Jime´nez et al., 1993, 1997; Wade et al., 1993; Vesonder et al., 1995) and capsicums (bell peppers) (Hashmi and Thrane, 1990; Utkhede and Mather, 2004). Other sources have been sorghum (Onyike and Nelson, 1992), sugar beets (Bosch and Mirocha, 1992), traditional African vegetables (van der Walt et al., 2006), tomatoes and lemons (Muniz et al., 2003), and black pepper (Pitt et al., 1998a). References. Booth (1971); Nelson et al. (1983); Desjardins (2006); Leslie and Summerell (2006).

5 Primary Keys and Miscellaneous Fungi

Fusarium verticillioides (Sacc.) Nirenberg Fig. 5.36 Fusarium moniliforme J. Sheld. (in part)

Teleomorph: Gibberella moniliformis Wineland Colonies on CYA usually covering the whole Petri dish, low at the margins to moderately deep centrally, of floccose to funiculose white mycelium; reverse pale, or in shades of pale salmon or violet. Colonies on MEA usually covering the whole Petri dish, low to moderately deep centrally, of white or pale salmon funiculose mycelium, sometimes powdery with microconidia; reverse of some isolates pale, of others violet or greyish magenta. Colonies

Fig. 5.36 Fusarium verticillioides (a) colonies on PDA and DCPA, 7 d, 258C; (b) phialides bearing chains of microconidia, bar ¼ 50 mm; (c) phialides, bar ¼ 10 mm; (d) macro- and microconidia, bar ¼ 10 mm

5.17 Genus Fusarium

on G25N 5–12 mm diam. At 58C, either germination or no growth. At 378C, colonies of 4–10 mm diam produced. On PDA, colonies of low, densely funiculose mycelium coloured white to pale salmon, usually powdery with chains of microconidia; reverse varying from isolate to isolate, pale salmon, greyish violet, brownish violet or deep violet; dark blue sclerotia produced by some isolates. On DCPA, colonies low, of thin, funiculose white to pale salmon mycelium in concentric rings, powdery with microconidia, and sometimes alternating with rings of macroconidia produced on the agar surface; reverse pale. Macroconidia usually long and slender, almost straight, thin walled, with foot-shaped basal cells; microconidia fusiform to clavate, produced from long monophialides, forming chains readily seen in situ under the low power microscope (6 to 10); chlamydoconidia not produced. Distinctive features. The production of microconidia in long chains from relatively long phialides distinguishes Fusarium verticillioides from most other common Fusarium species. F. proliferatum is very similar to in appearance to F. verticillioides but is distinguished from it by the production of polyphialides bearing microconidia in short chains. Taxonomy. In 1877, Saccardo described Oospora verticillioides from maize in Italy. In 1904, John Sheldon described Fusarium moniliforme from maize that was associated with animal toxicosis in the United States (see Desjardins, 2006), and for most of the twentieth century, the epithet F. moniliforme was used for this fungus. From an extensive study of the group of Fusarium species which included F. moniliforme, Nirenberg (1976) concluded that this species should be called F. verticillioides, but at the time, this proposal was not generally accepted. However, Seifert et al. (2003), representing the International Society for Plant Pathology and the International Committee on the Taxonomy of Fungi (ISPP/ICTF) Subcommittee on Fusarium Systematics, proposed that the name F. verticillioides be applied to the fumonisin producing fungus on maize, on the grounds that the name F. moniliforme represented an unacceptably broad species concept, and that F. verticillioides was undisputably the older name, which therefore had priority. This name is now generally accepted for isolates

121

from maize and a few other sources (Leslie and Summerell, 2006). Morphologically similar species within the Gibberella fujikeroi complex, associated with different plant hosts and different mating types, have been described: F. thapsinum from sorghum, F. sacchari from sugar cane, F. mangiferae from mango and F. fujikuroi from rice, and these names have now been taken up by Leslie and Summerell (2006). Physiology. The maximum temperature for growth of Fusarium verticillioides has been reported to be 32–378C, the minimum as 2.5–58C, and the optimum near 258C (Nirenberg, 1976). The minimum aw for growth was 0.87 at 258C, after a 4-month germination time (Armolik and Dickson, 1956). Reduction of headspace oxygen from 20 to 2% had no effect on growth of F. verticillioides, but vacuum packaging with oxygen absorbing sachets completely inhibited growth (Samapundo et al., 2007b). Fumonisins B1 and B2 were produced down to at least 0.92 aw (Marı´ n et al., 1995) but were not detected at 0.89–0.91 aw on irradiated maize grains (Marı´ n et al., 1999). Maximum fumonisin production occurred with 15% oxygen at 0.976 aw, but with 5% O2, 0.93 was the optimum aw (Samapundo et al., 2007b). Mycotoxins. The major mycotoxin produced by Fusarium verticillioides is fumonisin B1, the cause of leucoencephalomalacia in horses, pulmonary oedema in pigs and liver cancer in rats. It is a possible cause of human oesophageal cancer (Gelderblom et al., 1988; Sydenham et al., 1990; Shephard et al., 2007) and neural tube defects (Desjardins, 2006). For a comprehensive account of fumonisins, see Marasas (2001) and Desjardins (2006). Isolates of F. verticillioides from maize from many countries have been shown to produce fumonisins B1 and B2 (Bezuidenhout et al., 1988; Laurent et al., 1989; and see Pitt and Hocking, 1997; Desjardins, 2006). This species also commonly produces fusarins, fusaric acid and naphthoquinones, but not beauvarin, moniliformin or fusiproliferin (see Pitt and Hocking, 1997; Desjardins, 2006; Leslie and Summerell, 2006). Reports of zearalenone, diacetoxyscirpenol and deoxynivalenol production by F. verticillioides were discounted by Nelson et al. (1983). Recently identified isolates of F. verticillioides from banana rots (Mirete et al., 2004; Moretti et al., 2004) produce moniliformin but not

122

fumonisins (Patin˜o et al., 2006). These isolates need further characterisation. Ecology. Fusarium verticillioides is widespread in the tropics and humid temperate areas of the world (Leslie and Summerell, 2006). This species is an endemic pathogen of maize, causing both stalk rot and cob rot which can be present at all stages of plant development. It has been isolated from maize kernels in every major location where it has been sought, including the United States, Canada, Mexico, Honduras, Ecuador (Pacin et al., 2002), Peru and Argentina; from Iran (Ghiasian et al., 2004; 2006), India, China, Nepal (Desjardins and Busman, 2006) and Taiwan; from Europe (Logrieco et al., 2002), and from Australia, South Africa and other countries on the African subcontinent (Adejumo et al., 2007; Bigirwa et al., 2007) (see also Pitt and Hocking, 1997; Desjardins, 2006; Leslie and Summerell, 2006). Incidence in individual maize kernels, where reported, has almost always been very high, e.g. 100% in Sardinia (Bottalico et al., 1995), 54% in mainland Italy (Logrieco et al., 1995), 97% in Thailand (Pitt et al., 1993), 92% in the Philippines and 73% in Indonesia (Pitt et al., 1998a). Fusarium verticillioides (as F. moniliforme in earlier literature) has been reported as a pathogen of other Graminae, particularly sorghum (Leslie et al., 2005), rice (Desjardins et al., 2000a) and millet (Leslie et al., 2005) (see also Pitt and Hocking, 1997; Desjardins, 2006). It has been reported from several types of nuts, i.e. hazelnuts, pecans, peanuts and kola nuts (see Pitt and Hocking, 1997). F. verticillioides has also been reported from oilseeds: sunflower, amaranth, and soybeans; also spices, including coriander, fenugreek, cardamom and pepper (see Pitt and Hocking, 1997). It has also been found in mung beans (Pitt et al., 1994), biltong (van der Riet, 1976) and cheeses (Northolt et al., 1980). Fusarium verticillioides (or F. moniliforme) has been reported to cause storage rot of oranges and other citrus (Snowdon, 1990), garlic (Gargi and Roy, 1988), yams (Ogundana, 1972), pineapples (Lim, 1983), bananas (Chorin and Rotem, 1961; Jime´nez et al., 1993; Vesonder et al., 1995), asparagus (Snowdon, 1991) and dry rot of passion fruit (Lutchmeah, 1993). The molecular and chemical profiles of isolates of F. verticillioides that are pathogenic on bananas have been examined. Whilst banana isolates belong

5 Primary Keys and Miscellaneous Fungi

to mating population A (G. moniliformis), they are genetically distinct from F. verticillioides isolates from maize, produce moniliformin and are unable to produce fumonisins (Moretti et al., 2004; Mirete et al., 2004; Patin˜o et al., 2006). In light of recent taxonomic changes, many of the above reports of Fusarium moniliforme occurrence may need to be verified. References. Nirenberg (1976); Domsch et al. (1980); Nelson et al. (1983) (both under F. moniliforme); Desjardins (2006); Leslie and Summerell (2006).

5.18 Genus Geotrichum Link: Fr. For a long time, the genus Geotrichum has been distinguished by its method of reproduction, which is exclusively the formation of arthroconidia from vegetative hyphae. Arthroconidia are cylindrical spores formed by septation of vegetative hyphae into short segments, which separate at maturity. Young colonies sometimes appear yeast-like on CYA, but soon spread rapidly. Recently, the definition of Geotrichum has been broadened to include some species which also produce blastospores (yeast-like cells), but these are not of interest here. The most recent taxonomy is that of de Hoog et al. (1986), in which 23 species are described, together with two teleomorphs, Dipodascus and Galactomyces. Only one species, Geotrichum candidum, is significant in foods (Fig. 5.37).

Geotrichum candidum Link: Fr.

Fig. 5.37

Oidium lactis Fresen. Oospora lactis (Fresen.) Sacc.

Teleomorph: Galactomyces geotrichum (E.E. Butler & L.J. Peterson) Redhead & Malloch Colonies on CYA of variable size, 20–45 mm diam, very low and quite sparse, plane, of white mycelium, often leathery and difficult to dissect with a needle; reverse pale. Colonies on MEA 50– 65 mm diam, similar to on CYA but of softer, yeast-like texture. On G25N, no growth to

5.18 Genus Geotrichum

123

Fig. 5.37 Geotrichum candidum (a) colonies on MEA, 7 d, 258C; (b) arthroconidia, bar ¼ 10 mm

germination. At 58C, no germination to colonies up to 4 mm diam, of dense white mycelium. At 378C, usually no growth, occasionally sparse colonies up to 10 mm diam formed. Conidiophores undifferentiated hyphae, at maturity fragmenting almost entirely to form arthroconidia; arthroconidia hyaline, cylindrical, sometimes developing rounded ends and thickened walls, commonly 5–8  2–5 mm, smooth walled. Distinctive features. See genus description. Taxonomy. Some isolates of Geotrichum candidum have been reported to form a teleomorph, Galactomyces geotrichum, which consists of single ascospores within solitary asci borne along fertile hyphae (de Hoog et al., 1986). Physiology. Geotrichum candidum is restricted to habitats of high water availability, its minimum aw for growth being 0.90 (Heintzeler, 1939). Growth occurred at 0.95 but not at 0.90 aw (Plaza et al., 2003). Optimal growth temperatures are 25–308C (Plaza et al., 2003), with maxima of 35–388C (Domsch et al., 1980). The conidia have a very low heat resistance, with a D value of 30–40 min at 528C (Beuchat, 1981b). Miller and Golding (1949) reported that G. candidum was able to grow at very low oxygen tensions, but not anaerobically.

Mycotoxins. This species is not known to produce any toxic compounds. Ecology. Geotrichum candidum is a significant pathogen of citrus fruits during postharvest storage, causing sour rots (Butler et al., 1965; Hall and Scott, 1977; Snowdon, 1990). It occurs in fruit weakened by over maturity and long storage. Lemons and grapefruit are particularly susceptible, but a variety of other fruit can also be affected (Butler, 1960). Initial infection is mainly through injuries. The primary control measure is preventative, and lies in choosing sound, young fruit for long term storage (Hall and Scott, 1977). As G. candidum grows poorly below 108C, cold storage can assist in control (Rippon, 1980, Plaza et al., 2003). Geotrichum candidum causes spoilage of tomatoes (Okoli and Erinle, 1989; Snowdon, 1991), dried capsicums (Atanda et al., 1990) and sapodillas (Kusum and Geeta, 1990) in tropical countries. A wide variety of vegetables, including carrots, cucumbers, onions, peas and potatoes are also susceptible (Snowdon, 1991). Geotrichum candidum has long been a problem in the canning and freezing industries. Known as ‘‘machinery mould’’ (Eisenberg and Cichowicz, 1977), it is a frequent contaminant of processing lines, and consequently of products such as frozen foods (Pitt and Hocking, 1997). Standard methods

124

have been established for estimating G. candidum in food processing machinery, by physical counting rather than microbiological techniques (Cichowicz and Eisenberg, 1974). Improvements in sanitation procedures have reduced this problem in recent years. Kure et al. (2004) concluded that air was the major source of G. candidum contamination in cheese factories in Norway. This mould is a very common problem in raw milk in Europe especially when it is used in the manufacture of soft, fresh cheeses such as quarg or Roblochon (see Pitt and Hocking, 1997; Lopandic et al., 2006). Contamination of surface-ripened cheeses such as Brie and Camembert can also be a problem (Pitt and Hocking, 1997). However, recent surveys indicate that G. candidum may play an important role in the ripening of many soft and semi-hard cheeses as well as contributing desirable cheese flavours (Boutrou and Gueguen, 2005; Boutrou et al., 2006; Ghosh et al., 2006). Geotrichum candidum has been isolated from a variety of other foods, including meats, raw and Parma hams, hard cheeses and traditional fermented foods, and Nigerian alcoholic beverages (see Pitt and Hocking, 1997). Low levels were found in maize, peanuts, sorghum, soybeans and black rice in Thailand (Pitt et al., 1993, 1994). It is of quite common occurrence in barley during malting (see Pitt and Hocking, 1997). References. von Arx (1977); de Hoog et al. (1986).

5.19 Genus Hyphopichia Arx and van der Walt Hyphopichia is a genus of yeast-like fungi, probably closely related to Endomyces, from which it differs by the production of hyphal fragments which produce yeast-like conidia by budding from spicules. There is a single species, H. burtonii. Colonies on CYA 22–25 mm diam and on MEA 25–30 mm diam, of low and sparse to moderately dense white to pale grey mycelium, sometimes centrally umbonate; reverse pale. Colonies on G25N 10–12 mm diam, similar to those on CYA. No germination at 58C. At 378C, colonies 5–10 mm diam, similar to at 258C; reverse pale.

5 Primary Keys and Miscellaneous Fungi

Hyphopichia burtonii (Boidin et al.) Arx & van der Walt Fig. 5.38 Pichia burtonii Boidin et al. Endomycopsis burtonii (Boidin et al.) Lodder & Kreger Endomycopsis chodatii Wick.

Anamorph: Candida chodatii (Nechitsche) Berkhout Cladosporium chodatii (Nechitsche) Sacc. Trichosporon behrendii Lodder & Kreger Trichosporon variabile (Lindner) Delitsch

Conidia borne from spicules (small projections) along the length of undifferentiated hyphae, yeastlike, ellipsoidal, pyriform or near cylindroidal, 2.5– 5.0  1.5–2.5 mm, with thin, smooth walls; hyphal fragments also evident, 10–50  2.5–3.0 mm, still functioning as fertile hyphae, producing conidia (blastospores) from lateral spicules and terminal buds. Ascospores not observed in pure culture, two mating strains needed; ascospores hat-shaped, 1–4 per ascus. Distinctive features. This species produces white, powdery, filamentous colonies which bear conidia from spicules on vegetative hyphae, and also by budding (yeast-like cells). It is distinguished from E. fibuliger by the fragmentation of hyphae into short lengths; these can also act as reproductive structures. Taxonomy. This species and Endomyces fibuliger have rarely been placed in the same genus by taxonomists, but in culture the resemblance is unmistakable. Both grow at more or less the same rates and have similar habitats. One (E. fibuliger) is homothallic and the other heterothallic; however, the fragmenting of fertile hyphae in Hyphopichia burtonii is the only obvious difference in morphology. The teleomorph of Hyphopichia burtonii has been seen only in the laboratory, after mixing of two mating strains. However, the teleomorph name is used here, both because it has become accepted in the recent literature and because there are several confusing anamorph names, at both genus and species level. Some recent taxonomies have included this species in the yeasts, as Pichia burtonii Boidin et al. (Kurtzman and Fell, 1998; Barnett et al., 2000). Physiology. Hyphopichia burtonii grows more rapidly at 308C than 208C (Ramakrishna et al., 1993). The minimum is >58C and growth occurs at 378C. Growth occurs at least down to 0.90 aw on barley grains (Ramakrishna et al., 1993). Mycotoxins. This type of fungus is unlikely to produce toxic compounds.

5.20 Genus Lasiodiplodia

125

Fig. 5.38 Hyphopichia burtonii (a) colonies on CYA and MEA, 7 d, 258C; (b) vegetative cells, bar = 10 mm; (c,d) vegetative cells, bar = 5 mm

Ecology. Hyphopichia burtonii is not uncommon in cereals and cereal products, especially packaged bread (Spicher, 1984a, 1985). In Europe, along with Endomyces fibuliger, it is known as ‘‘chalky mould’’ (Spicher, 1986b). It is also part of the flora of Parma hams during ripening (Simoncini et al., 2007). Reference. Barnett et al. (1990).

characterised by distinctive large, striate conidia, as described below.

Lasiodiplodia theobromae (Pat.) Griffon & Maubl. Fig. 5.39 Botryodiplodia theobromae Pat.

Teleomorph: & Curt.) Arx

Botryosphaeria

rhodina

(Berk.

5.20 Genus Lasiodiplodia Ellis and Everh. Colonies on CYA and MEA usually filling the Lasiodiplodia is the correct generic name for the fungus previously known as Botryodiplodia theobromae Pat. (Sutton, 1980). Both genus and species are

whole Petri dish with a loose to moderately dense weft of light to dark grey hyphae, adhering to the Petri dish lid; pycnidia borne beneath the agar surface (visible from the reverse), and sometimes

126

5 Primary Keys and Miscellaneous Fungi

Fig. 5.39 Lasiodiplodia theobromae (a) colonies on CYA, 7 d, 258C; (b) conidia, bar = 10 mm

at the base–lid interface or on the lid itself; reverse pale, black near pycnidia, uniformly grey-black elsewhere. On G25N, colonies 30–50 mm diam, occasionally more, low and spreading, with sparse white aerial mycelium; reverse pale or grey. No growth at 58C. At 378C, colonies greater than 50 mm diam, of low, dense mycelium, coloured grey or deep red; red brown soluble pigment and occasionally exudate produced; reverse deep red brown to almost black. Reproductive bodies on CYA pycnidia, greyblack and roughly spherical, 200–400 mm diam, but on other media reportedly up to 5 mm diam (Punithalingam, 1976), or forming a stroma (Alasoadura, 1970), easily ruptured, of dark brown to black pseudoparenchymatous cells; conidia at 7 days ellipsoidal, 15–20  9–12 mm, with smooth, brownish walls and little ornamentation, at maturity (18–)20–30  10–15 mm, with a transverse septum and ornamented by longitudinal striations. Distinctive features. Lasiodiplodia theobromae is distinguished in culture by fast growth on G25N and at 378C: on G25N growth is sparse and usually colourless, but at 378C it is dense, with mycelium grey and/or red. Microscopically, this species is distinguished by large ellipsoidal conidia which are borne in black pycnidia, and are ornamented with a transverse septum and longitudinal striations.

Taxonomy. A molecular study using a combined analysis of the ITS region and the translation elongation factor 1-a indicated that Lasiodiplodia theobromae as currently understood includes three species, distinguishable by differences in the size of their conidia (Alves et al., 2008). This species is sometimes known by its teleomorph name, Botryosphaeria rhodina, but this state is not usually seen in culture. Physiology. Alasoadura (1970) reported cardinal temperatures for Lasiodiplodia theobromae as minimum 158C, optimum 288C and maximum 408C. This is difficult to reconcile with statements by Uduebo (1974) that ‘‘The vegetative growth at 378C is scanty’’, or that this species does not grow at 358C on potato dextrose agar (Adeniji, 1970b). Isolates of this species which we have studied grew rapidly at 378C, and our observations are consistent with the statement that distribution of L. theobromae is mainly confined to the area between 408N and 408S (Punithalingam, 1976). Several references suggest that germination and growth of L. theobromae is confined to high water activities: vigorous growth observed on G25N in this study is at variance with this. Mycotoxins. Mycotoxin production has not been reported. Ecology. Punithalingam (1976) aptly described Lasiodiplodia theobromae as ‘‘an unspecialised virulent rot pathogen causing numerous diseases:

5.21 Genus Monascus

dieback, root rot, rot or decay of various fruits and storage rot of yams.’’ Spoilage by Lasiodiplodia of avocados, bananas, grapes, guavas, mangoes, okra, papayas, passionfruit, traditional Nigerian foods and yams has been reported (see Pitt and Hocking, 1997). Control measures for fruits are well documented by Punithalingam (1976). Lasiodiplodia theobromae has also been reported from pecans, peanuts, wheat and soybeans (see Pitt and Hocking, 1997). In our experience, it is of common occurrence in many tropical food commodities. We isolated it from 58% of Thai maize samples, at up to 46% of kernels in infected samples and in 7% of all kernels examined (Pitt et al., 1993). It was also common in Thai peanuts, mung beans and sorghum, with 30–33% of samples showing infection (Pitt et al., 1993, 1994). Indonesian crops were equally affected, particularly peanuts, where 47% of samples were positive, with up to 64% of kernels affected in those samples, and overall 9% of the more than 12,000 peanut kernels examined were infected with this species. More than 25% of maize, talo bean, velvet bean, black soybean and cowpea samples also showed infection (Pitt et al., 1998a). References. Punithalingam (1976); Domsch et al. (1980), under Botryosphaeria.

5.21 Genus Monascus Tiegh. Monascus is a genus of Ascomycetes characterised by the production of colourless to pale brown cleistothecia and aleurioconidia. Each cleistothecium is borne from a knot of hyphae on a well defined stalk, in 7 day old cultures resembling a clenched fist on a narrow forearm. Aleurioconidia occur singly or in short chains. Asci break down rapidly so that, when ascospores are mature, the impression under the microscope is of a sac filled with a mass of ellipsoidal, smooth walled, refractile spores. Species of Monascus are best known for their role in the fermentative production of Oriental foods, of which red rice (ang-kak), rice wine and kaoliang brandy are the best known (Hesseltine, 1965; Lin, 1975). Three species were accepted in the most recent revision of the genus (Hawksworth and Pitt, 1983): two, M. purpureus Went and M. pilosus K. Saito ex D. Hawksw. & Pitt, are associated almost exclusively

127

with fermented foods; the third, M. ruber Tiegh., is uncommon in fermented foods but of widespread occurrence elsewhere, and it sometimes causes food spoilage. Only M. ruber is treated here. For information on the other species, see Hawksworth and Pitt (1983). Further species have been described from Iraqi soils (Cannon et al., 1995) but these are not relevant to foods.

Monascus ruber Tiegh.

Fig. 5.40

Colonies on CYA 20–32 mm diam, occasionally only 15 mm, plane, sparse, surface texture floccose to deeply floccose; mycelium white at first, then becoming pale brown as cleistothecia and aleurioconidia develop, in age sometimes becoming dark brown; brown soluble pigment sometimes produced; reverse sometimes uncoloured, usually brown to dark brown near sepia. Colonies on MEA 30–42 mm diam, plane and sparse, sometimes with little aerial growth but usually with deeply floccose mycelium, white at the margins, becoming brown to orange brown as cleistothecia and aleurioconidia develop; orange or brown soluble pigment sometimes produced; reverse pale at the margins, khaki or brownish orange at the centres. Colonies on G25N usually 16–20 mm diam, sometimes only 10 mm, plane, sparse and floccose; mycelium white; reverse uncoloured or brown. No growth at 58C. At 378C, colonies 12–30 mm diam, low and sparse, coloured as on CYA at 258C, or with reverse deeper brown and sometimes with a reddish tinge. Cleistothecia spherical, 30–60 mm diam, borne as a hyphal knot from a well-defined stalk, with cellular walls, becoming brown during maturation; ascospores ellipsoidal, hyaline, 5–7  4.0–4.5 mm, smooth walled. Aleurioconidia sometimes borne on pedicels from the sides of hyphae, but more commonly terminally, sometimes borne singly but more often in chains of up to 10 cells long, spherical to pyriform, often rounding at maturity, 10–14 mm diam or 10–18  8–14 mm, with thick, smooth, brown walls. Chlamydoconidia and arthroconidia produced by most isolates also. Distinctive features. The cleistothecium produced by Monascus is distinctive, being borne as a fist-like hyphal knot on an arm-like stalk. M. ruber is distinguished from other Monascus species by relatively rapid growth, especially on MEA, and by brown pigmentation in the walls of cleistothecia and aleurioconidia.

128

5 Primary Keys and Miscellaneous Fungi

Fig. 5.40 Monascus ruber (a) colonies on CYA and MEA, 7 d, 258C; (b) cleistothecia, bar = 10 mm; (c) aleurioconidia, bar = 10 mm; (d) ascospores, bar = 5 mm

Taxonomy. Sequencing of the ITS region and part of the b-tubulin gene indicate that Monascus ruber and M. pilosus should be considered as a single species (Park et al., 2004a). Physiology. The cardinal temperatures reported for Monascus ruber are minimum 15–188C; optimum 358C and maximum near 458C (Manandhar and Apinis, 1971; Panagou et al., 2003). M. ruber has been isolated from relatively concentrated substrates (such as high moisture prunes, ca. 0.85 aw) and is probably weakly xerophilic. However, an isolate failed to grow

in a solution of NaCl at 0.92 aw (Panagou et al., 2003). Growth of M. ruber was little affected over the pH range 3.0–5.0 (Panagou et al., 2003) and was able to grow down to pH 2.2 at 308C in high aw (Panagou et al., 2005). The combined effect of temperature (25–358C), water activity (0.999–0.92 aw) and pH (2–6.8) was studied on gradient plates by Panagou et al. (2005), and also modelled against several equations (Panagou et al., 2003, 2005, 2007). Ascospores of Monascus ruber are heat resistant. D80 in citrate buffer (pH 4) was 2.1 min. NaCl

5.22 Genus Moniliella

129

provided some protection: D80 values were 0.88 min in 5.6% brine, but 1.04 min in 10.5% brine. z values over the temperature range 70–808C varied from 7.4 to 7.9 C8 under the conditions used (Panagou et al., 2002). They concluded that a heat process of 5 min at 808C would assist the olive industry in producing a stable product. Pigmentation has received a great deal of attention, having been used to distinguish a number of species utilised in Oriental fermentations. However, Carels and Shepherd (1977) showed that red pigments form during growth near neutral pH values, whereas cultures become orange if the pH of the fermenting food becomes strongly acid. Pigment stability is affected by pH, temperature, light, oxygen and water activity (Dufosse et al., 2005). The production of pigments by Monascus species has been reviewed by Miyake et al. (2008). Mycotoxins. Monascus ruber (and M. purpureus) produce citrinin (Blanc et al., 1995a, b) both in submerged and solid state culture. This is of importance as red pigments produced by Monascus species are used as food colourants. Recent studies indicate that high pigment production can be achieved without citrinin synthesis by incorporating histidine in the growth media (Hajjaj et al., 2000; Xu et al., 2003). Wang et al. (2005) surveyed type cultures of 23 species of Monascus (most now regarded as synonyms) for their ability to synthesise citrinin and found that all, including M. ruber, produced citrinin. Ecology. A widespread species, Monascus ruber has caused spoilage of high moisture Australian

prunes (Hawksworth and Pitt, 1983). It was isolated quite frequently from Indonesian dried fish (Wheeler et al., 1986). Other sources include mayonnaise (Muys et al., 1966a), bread (Spicher and Isfort, 1988), table olives (Panagou et al., 2002), and minced meat, soft cheese, fruit sauce, spices, honey, cacao beans, soy beans, peas, palm kernels, maize and various animal feeds, and silage (data sheets, International Mycological Institute, Egham, Surrey; Hawksworth and Pitt, 1983). References. Domsch et al. (1980); Hawksworth and Pitt (1983).

5.22 Genus Moniliella Stolk and Dakin Moniliella is a genus of yeast-like fungi, characterised by the production of budding cells, thin walled arthroconidia and, in one species, by relatively large chlamydoconidia (up to 3 hyphal diameter). In some respects it is similar to Hyphopichia, but conidia are larger, and the hyphal fragments bearing conidia on spicules, characteristic of Hyphopichia, are absent. The latest taxonomy (de Hoog, 1979) accepts two species, M. suaveolens and M. acetoabutans. M. suaveolens is found in butter and margarine, and other substrates rich in oil, while M. acetoabutans is of interest because of its resistance to acetic acid and hence potential ability to spoil mayonnaise and other products preserved with acetic acid.

Key to Moniliella species included here 1

Spherical chlamydoconidia produced, 8–12 mm diam, with thick, brown walls; growth on malt acetic agar Thick walled chlamydoconidia not produced; no growth on malt acetic agar

Moniliella acetoabutans Stolk & Dakin Fig. 5.41 Colonies on CYA and MEA 22–30 mm diam, deeply floccose, especially on CYA, mycelium pure white; reverse yellow brown. Colonies on G25N 5–10 mm diam, deep but often mucoid, uncoloured. No growth at 58C. Usually no growth at 378C, occasionally colonies up to 5 mm diam. Conidia of three types produced, budding cells from hyphal extremities, arthroconidia by differentiation of

M. acetoabutans M. suaveolens

hyphal tips, and chlamydoconidia, in intercalary or terminal positions on hyphae, solitary or in short chains; budding conidia ellipsoidal, arthroconidia cylindroidal, both 5–9 mm long, chlamydoconidia spherical, 8–12 mm diam, with thick brown walls. Distinctive features. Moniliella acetoabutans is morphologically distinguished by its three types of conidia: budding conidia like a hyaline Cladosporium; arthroconidia like Geotrichum, but not so extensive; and relatively large, brown walled chlamydoconidia. In culture, the definitive test for M.

130

5 Primary Keys and Miscellaneous Fungi

Fig. 5.41 Moniliella acetoabutans (a) colonies on CYA and MEA, 7 d, 258C; (b,c) conidiophores producing budding cells (slide culture), bar = 10 mm; (d) arthroconidia, bar = 10 mm; (e) chlamydoconidia, bar = 10 mm

acetoabutans is its ability to produce quite rapidly growing white colonies on malt acetic agar – or even on MEA containing 2% acetic acid. Physiology. This species appears to be unique in its tolerance of acetic acid and other weak acid preservatives. In our laboratory, Moniliella acetoabutans has been able to grow in MEA with 4% added acetic acid, to our knowledge a unique property. This species is capable of fermentative growth, like a true yeast, and also appears to be highly tolerant of acid pH. Mycotoxins. Toxic compounds are not produced.

Ecology. Dakin (Stolk and Dakin, 1966) isolated Moniliella acetoabutans from spoiling sweet fruit sauce, and then a variety of other similar products. It also caused fermentative spoilage of a large production run of mayonnaise in Australia in 1971. The source was wooden vinegar tanks, where the fungus was surviving in 10% acetic acid, then growing on the wood as the tank level lowered and the acetic acid became diluted by evaporation. Data sheets from the International Mycological Institute, Egham, Surrey, record isolations from various acetic acid preserves.

5.23 Genus Nigrospora

131

Moniliella acetoabutans infections fortunately appear to be rare. They may be overcome either by holding vinegar or acetic acid in stainless steel tanks, or by pasteurising this ingredient before addition to product. References. Stolk and Dakin (1966); Dakin and Stolk (1968); de Hoog (1979).

Moniliella suaveolens (Lindner) Arx

Fig. 5.42

Cladosporium suaveolens (Lindner) Delitsch Cladosporium butyri O.S. Jensen

Colonies on CYA and MEA highly variable in character, 15–40 mm diam, either white, floccose, sparse and persistently white, or low, dense, velutinous and olive, lightly to heavily sporing; reverse colourless or olive. Colonies on G25N 3–8 mm diam, of white mycelium. At 58C, sometimes germination. At 378C, no growth. Conidiophores undifferentiated, bearing conidia in short, sometimes branched, acropetal chains (the youngest spore at the end), breaking up in liquid mounts; conidia subspheroidal or ellipsoidal to cylindroidal, or somewhat irregular, nonseptate, when ellipsoidal 9–13  7–10 mm, when cylindroidal commonly 15–20  5–7 mm, with smooth, slightly thickened walls. Distinctive features. Unlike Moniliella acetoabutans, this species does not make chlamydoconidia. It is not preservative resistant, and is normally associated with oils and oil-based foods.

Taxonomy. The taxonomy of Moniliella suaveolens is complicated by the fact that the two varieties accepted by de Hoog (1979) culturally bear little resemblance to each other. One (M. suaveolens var. suaveolens) produces low, spreading colonies, which remain white; the other (M. suaveolens var. nigra) forms compact, dense colonies, which rapidly become olive. Their incorporation into a single species by de Hoog (1979) was due to similarities in conidial production and conidia. Physiology. No physiological studies are known to us. Mycotoxins. Mycotoxins are not produced. Ecology. This species has caused spoilage of margarine in Europe (Muys et al., 1966a, b; Bours and Mossel, 1973) and Australia (Hocking and Pitt, unpublished). It has been reported as one of the causes of chalky bread in Europe (Spicher, 1985, 1986b; Spicher and Isfort, 1987). It has occasionally been isolated from cheese (Chabalier et al., 1997; Torkar and Vengust, 2008). Reference. de Hoog (1979).

5.23 Genus Nigrospora Zimm. Characterised by the production of relatively large, solitary, jet black, smooth-walled, oblate conidia, Nigrospora occurs in nature mainly as a plant and seed pathogen, but is found also in air, soil and

Fig. 5.42 Moniliella suaveolens (a) colonies on MEA, 7 d, 258C; (b,c) vegetative cells (slide culture), bar = 10 mm

132

water. Two species are of significance in foods: Nigrospora oryzae and Nigrospora spherica.

Nigrospora oryzae (Berk. & Broome) Petch Fig. 5.43 Teleomorph: Khuskia oryzae H.J. Huds. Colonies on CYA and MEA covering the whole Petri dish, low to moderately deep, dense to floccose, mycelium flesh coloured or pale orange to pure grey; black conidia conspicuous at low magnifications; reverse pale, greyish orange or grey to deep bluish grey. Colonies on G25N usually 10– 15 mm diam, white or mucoid. No growth at 5 or 378C. Conidiophores borne from aerial hyphae, short, dark walled, bearing conidia in isolation or in clusters from groups of irregular cells; conidia solitary, jet black, oblate, sometimes collapsing, mostly 12–15 mm long, with smooth, featureless walls, remaining attached or (on natural substrates) violently discharged. Distinctive features. See genus description. Nigrospora is clearly distinguished from Arthrinium, which it superficially resembles, by its jet black conidia which lack surface features or markings.

5 Primary Keys and Miscellaneous Fungi

Taxonomy. Nigrospora oryzae produces an ascomycete teleomorph, Khuskia oryzae, which is found only as a pathogen on certain plants. Physiology. Optimum growth for Nigrospora oryzae has been recorded at 0.995 aw and 258C. N. oryzae sporulated at 0.98 and 0.995 aw but not at lower aw values (0.95, 0.90 or 0.85) at 258C (Sempere and Santamarina, 2006). Inability to grow at low or high temperatures has been noted above. Mycotoxins. Mycotoxin production has not been reported for Nigrospora species. Ecology. As a plant pathogen on cereal crops, the widespread distribution of Nigrospora oryzae in cereal grains is to be expected. It has been reported from barley (Fakhrunnisa et al., 2006), wheat, other leguminous and cereal foods, pecans and various health foods (see Pitt and Hocking, 1997). It also causes a storage disease of apples in India (Khanna and Chandra, 1975). We isolated Nigrospora oryzae frequently in surveys of Southeast Asian food commodities (Pitt et al., 1993, 1994, 1998a and unpublished). The highest level of occurrence was on paddy rice, where it was isolated from an average of 40% of samples, with up to 68% of infected kernels in infected samples and in 10% of more than 7,000 grains examined from Thailand, Indonesia and the Philippines. This species was isolated from high percentages of samples (25% or more) of maize, sorghum, soybeans, cashews and copra from Thailand, with overall infection rates of 2–5%. In Indonesia,

Fig. 5.43 Nigrospora oryzae (a) colonies on MEA, 7 d, 258C; (b) conidia, bar = 10 mm

5.24 Genus Pestalotiopsis

kemiri nuts showed infection at 5% overall, while maize, sorghum, soybeans, kidney beans, cowpeas and talo beans were all infected at the rate of 1% or more of all seeds examined (Pitt et al., 1998a). Maize, mung beans and black pepper from the Philippines showed similar levels of infection (our unpublished data). Additional species. Nigrospora spherica (Sacc.) E.W. Mason is similar in culture to N. oryzae, but produces larger conidia, mostly 15–18 mm long. This species is associated with two diseases of bananas, crown rot (Wallbridge, 1981) and squirter (Hall and Scott, 1977; Snowdon, 1990), and Nigrospora rot of apples (Snowdon, 1990). It has also been recorded from wheat, peanuts, pecans, maize and biltong (see Pitt and Hocking, 1997). Although less common in Southeast Asian food commodities than Nigrospora oryzae, we isolated N. spherica from kemiri nuts, maize, peanuts, milled rice, cowpeas and black soybeans in Indonesia (Pitt et al., 1998a), and from paddy rice, maize, soybeans and black pepper in the Philippines (our unpublished data). References. Ellis (1971); Domsch et al. (1980).

5.24 Genus Pestalotiopsis Steyaert Pestalotiopsis and the closely related genus Truncatella Steyaert (see below) are characterised by the formation of black acervuli containing relatively large fusiform

133

conidia with three or four transverse septa and spiky appendages from one or both ends. Neither genus is encountered frequently in foods other than cereals or nuts, where they can be spoilage agents. In our experience, direct plating of cereals will frequently detect Pestalotiopsis, but dilution plating techniques are ineffective. The name Pestalotia De Not. has frequently been incorrectly applied to foodborne fungi which should have been identified as Pestalotiopsis or, less frequently, Truncatella. In his comprehensive revision, Steyaert (1949) confined Pestalotia to a single species, not found in foods. Sutton (1980) accepted a single species in Pestalotiopsis, P. guepinii. However, many new species have been described since, often based on host specificity. Molecular studies on ITS and 5.8S DNA have shown that host specificity is not a sound basis for species delimitation in Pestalotiopsis (Jeewon et al., 2004).

Pestalotiopsis guepinii (Desm.) Steyeart Fig. 5.44 Colonies on CYA and MEA growing rapidly, covering the whole Petri dish, plane and floccose; mycelium usually white, sometimes off-white to pale brown; reverse pale or in similar colours to the mycelium. Colonies on G25N 10–16 mm diam, of low, white mycelium. Sometimes germination or growth at 58C. Usually no growth at 378C.

Fig. 5.44 Pestalotiopsis guepinii (a) colonies on CYA, 7 d, 258C; (b) conidia, bar = 10 mm

134

Conidia produced in flat, black acervuli, borne just beneath the agar surface, opening irregularly at maturity, filled with a dense layer of conidia; conidia fusiform, five celled (four septate), 20–28  6–9 mm, the central 3 cells brown, 15–20 mm long, the apical and basal cells hyaline, the basal one with a single usually unbranched spike-like appendage and the apical one with two or more simple or branched spiky appendages. Distinctive features. Pestalotiopsis shares with Truncatella the production in subsurface acervuli of relatively large fusiform conidia with appendages. In Pestalotiopsis, conidia have four septa, in Truncatella, three. Physiology. Optimum sporulation of Pestalotiopsis guepinii was achieved on media containing 3% sucrose. In addition, reduced nitrogen concentration also favoured sporulation (Ebenezer et al., 2002). Mycotoxins. Mycotoxin production has not been reported. Ecology. There are few records under the name Pestalotiopsis in the food literature. Most relate to fruit spoilage: Pestalotiopsis sp. from crown rot of bananas and rots in litchis, canker in guavas, and tarry rots in pomegranates (Snowdon, 1990). Most reports of Pestalotia species, which probably refer to Pestalotiopsis, have come from pecans. Other Pestalotia records include wheat, rice, almonds and hazelnuts (see Pitt and Hocking, 1997). We isolated Pestalotiopsis guepinii at low levels from Thai copra, maize, black beans and red beans (Pitt et al., 1993, 1994). Additional genus. Truncatella Steyaert produces colonies similar to those of Pestalotiopsis, and produces similar conidiomata. Conidia are fusiform with three septa, and measure 15–20  6–8 mm; the two median cells are brown and 11–14 mm long; apical and basal cells are hyaline; and the basal cell is without appendages. Appendages from the apical cell are variable in number and branching. No records of Truncatella in foods were found, but it is probable that some references to Pestalotia include Truncatella species. Reference. Sutton (1980).

5.25 Genus Phoma Sacc. Phoma is a large and variable genus, characterised by the production of small, single celled conidia in pycnidia. The pycnidia in Phoma are black, have

5 Primary Keys and Miscellaneous Fungi

one or more small ostioles (orifices), are produced beneath the agar surface, and exude the conidia in slime. Several Phoma species have been reported from foods, but until recently the taxonomy of the genus was uncertain, so identifications to species were often unreliable. A new monograph has clarified the taxonomy of the genus (Boerema et al., 2004). This book provides keys and descriptions of 223 species and subspecies in 10 sections, with a great deal of other information including methods for identifying species in natural habitats. A single species, Phoma sorghina, is described here as representative of the genus.

Phoma sorghina (Sacc.) Boerema

Fig. 5.45

Colonies on CYA and MEA variable, usually 50–55 mm diam, of dense to floccose grey green or olive mycelium, with characteristic white to salmon pink tinges; reverse salmon or reddish. Colonies on G25N 8–10 mm diam, of sparse brown mycelium. No growth at 5 or 378C. Pycnidia produced abundantly on MEA, just beneath the agar surface, 300–400 mm diam, with one or more inconspicuous ostioles exuding conidia in a slimy matrix; conidia cylindroidal, 4–5  2–2.5 mm, hyaline, with thin, smooth walls. Distinctive features. Phoma species produce black subsurface pycnidia which exude small cylindrical conidia in slime. These features are sufficient in the present context; however, see Sutton (1980) for a range of other similar genera. Taxonomy. Taxonomic problems in Phoma have been addressed by Kovics et al. (2005) and a molecular taxonomy has recently been constructed (Aveskamp et al., 2006) for the 223 species and subspecies accepted by Boerema et al. (2004). Physiology. Sporulation by Phoma species may be stimulated by growth on DCPA or PDA. Carnation Leaf Agar, developed for Fusarium species, induces sporulation in most Phoma isolates. Mycotoxins. Phoma sorghina has been reported to produce tenuazonic acid (Shephard et al., 1991) and P. lingam the first naturally occurring epimonothiodiocoperazine, named sirodesmin H (Pedras et al., 1988). No studies have been reported on the significance, if any, of production of these toxins by these species.

5.25 Genus Phoma

135

Fig. 5.45 Phoma sorghina (a) colonies on CYA and MEA, 7 d, 258C; (b) pycnidium, bar = 10 mm; (c) conidia, bar = 5 mm

Ecology. Phoma sorghina is the principal Phoma species found in grain sorghum. Other Phomas, usually unidentified to species, have been reported from cereals, including barley, wheat, maize and rice (see Pitt and Hocking, 1997; Do Amaral et al., 2005; Perello and Moreno, 2005). Phomas, sometimes identified to species, have been reported from a wide variety of fruits and vegetables (Snowdon, 1990, 1991). Important are black rot of papayas due to P. caricae-papayae (Tarr) Punith., black rot of melons due to Phoma cucurbitacearum (Fr.) Sacc. (Snowdon, 1990), crown rot of bananas caused by P. sorghina and Phoma rot of sugar beets due to P. betae A.B. Frank. Other records have come from pecans, peanuts, apples, potatoes, fresh vegetables,

raw milk, spoiled cheese, sunflower seed, coriander seed, infant cereal food, biltong and a variety of Japanese foods (see Pitt and Hocking, 1997). Phoma species (unidentified) were common in paddy rice from Thailand (in 56% of samples, up to 44% of grains in infected samples and 8% of all grains examined) (Pitt et al., 1994). Similar levels occurred in sorghum from the Philippines and black soybeans from Indonesia (Pitt et al., 1998a and our unpublished data). Lower but significant levels (1–2% infection overall) were found in sorghum, mung beans and soybeans from Thailand, paddy rice and soybeans from the Philippines and paddy rice, mung beans, soybeans and talo beans from Indonesia (Pitt et al., 1994, 1998a and our unpublished data).

136

5 Primary Keys and Miscellaneous Fungi

Although they occur in a variety of foods, Phoma species are most likely to cause spoilage in weatherdamaged cereals. References. Boerema et al. (2004).

5.26 Genus Stemphylium Wallr. Like Alternaria and Ulocladium, Stemphylium produces conidia with both longitudinal and transverse septa. Stemphylium conidia are more or less ellipsoidal to short cylindroidal, like those of Ulocladium. Whereas those of Ulocladium are narrower at the base than the apex, and can be pointed at one end, to give a pyriform or apiculate shape, those of Stemphylium have rounded ends and are usually symmetrical longitudinally. For further details concerning the differences among these three genera, see the section dealing with Alternaria in this chapter, or Simmons (1967). In an extensive molecular study of Alternaria, Stemphylium and related genera (Ulocladium, Embellisia and Nimbya), Pryor and Gilbertson (2000) and Pryor and Bigelow (2003) provided strong molecular evidence that the genus Stemphylium is distinct from these other genera. Species of Stemphylium occur commonly as weak parasites or saprophytes on a variety of plants and plant materials. The taxonomy of the genus has recently been revised: molecular methods showed good agreement with existing morphological taxonomy (Caˆmara et al., 2002). One species is treated here, S. botryosum (Fig. 5.46).

Stemphylium botryosum Wallr.

Fig. 5.46

Teleomorph: Pleospora tarda E.G. Simmons Colonies on CYA 50–65 mm diam, of low to somewhat floccose, rather sparse mycelium, olive to olive brown; reverse olive grey to dark grey. Colonies on MEA 50–70 mm diam, low to floccose, in the latter case sometimes with areas of white or pale yellow mycelium, elsewhere pale to dark grey; reverse light to dark grey, sometimes pale yellow or pinkish. Colonies on G25N 6–15 mm diam, pale to dark grey. At 58C, usually germination. At 378C, no growth. Colonies on DCMA 35–45 mm diam, low to somewhat floccose, sparse, grey to dark grey in areas; reverse in similar colours.

Conidiophores undifferentiated, of variable length, with slightly swollen (up to 7 mm) tips bearing conidia successively, blown out from a pore as a hyaline cell, then septating both longitudinally and transversely, becoming thickwalled and detaching with maturity; the smaller conidia more or less spherical, with one to two septa, the larger ones at maturity usually short cylindrical, with rounded ends and straight or irregularly swollen sides, with two to four transverse and one to two longitudinal septa, sometimes near spherical or irregular in shape, overall ranging in size from 13 mm diam up to 30–32  16–20 mm, with brown walls, slightly roughened or with short spines, or occasionally with larger dark warts. Distinctive features. As noted above, Stemphylium is distinguished by dark conidia with both transverse and longitudinal septa, with ends rounded, symmetrical from end to end. S. botryosum conidia characteristically are slightly constricted in the middle. Taxonomy. The teleomorphs of this and other Stemphylium species are in the Ascomycete genus Pleospora (Simmons, 1969, 1985). They are not often seen in pure culture in the Petri dish. Metabolite profiles have been used to distinguish several Stemphylium species, including S. botryosum (Andersen et al., 1995). Physiology. Potato dextrose agar supported abundant mycelial growth, whereas conidiogenesis was highest on Sabouraud maltose agar (Susuri and Doga-Gashi, 2003). Mycotoxins. This species has been reported to produce a toxin called stemphol, but mammalian toxicity was not indicated (Solfrizzo et al., 1994). Ecology. This and other Stemphylium species have been reported as pathogens on mangoes (Johnson et al., 1990), squash (Assawah and Zarari, 1984), sweet potatoes (Ravichandran and Sullia, 1983), asparagus and lettuce (Snowdon, 1991), tomatoes (Dodds et al., 1991; Muniz et al., 2003) and peppers (Muniz et al., 2003). Stemphylium species have also been isolated from malting barley (Sepitkova and Jesenska, 1986), lucerne seeds (Susuri and Doga-Gashi, 2003) and soybeans (our unpublished data). References. Simmons (1967, 1969, 1985).

5.27 Genus Trichoconiella

137

Fig. 5.46 Stemphylium botryosum (a) colonies on CYA and MEA, 7 d, 258C; (b) conidiophores and conidia in situ, bar = 50 mm; (c,d) conidia, bar = 10 mm

5.27 Genus Trichoconiella B.L. Jain This genus was established to accommodate the fungus more commonly known as Alternaria padwickii (Ganguly) M.B. Ellis. Conidia of this species produce only transverse septa, excluding it from Alternaria (Jain, 1975).

Trichoconiella padwickii (Ganguly) B.L. Jain Fig. 5.47 Alternaria padwickii (Ganguly) M.B. Ellis Colonies on CYA 30–45 mm diam, plane, dense to floccose, mycelium yellowish grey to light orange,

sometimes with pale grey areas; yellow exudate sometimes produced; reverse pale or orange at the margins, centrally very dark brown to bluish black. Colonies on MEA 45–55 mm diam, plane, low to floccose, mycelium pale orange or grey; reverse pale or pinkish at the margins, overall bluish black. No growth on G25N or at 58C. At 378C, colonies up to 5 mm diam formed, or no growth. Colonies on DCMA 35–55 mm diam, similar in appearance to on MEA, but mycelium mid to dark grey; reverse grey, dark grey or bluish black. Conidia best observed on colonies grown on sterile wheat grains, borne singly from long hyphae, with five to six transverse septa, 40–55  9–11 mm between the tip and last septum, with the central

138

5 Primary Keys and Miscellaneous Fungi

Fig. 5.47 Trichoconiella padwickii (a) colonies on CYA and MEA, 7 d, 258C; (b,c) conidia, bar = 10 mm

cells larger, giving a narrowly ellipsoidal shape, and with smooth, brown walls; hyphae sometimes remaining attached to the conidium and then giving the impression of long appendages. Distinctive features. This genus superficially resembles Alternaria, but produces conidia with transverse septa only. Long, whip-like hyphal appendages are also distinctive. Trichoconiella padwickii appears to be almost exclusively associated with rice.

Physiology. No reports of physiological studies are known to us. Mycotoxins. Mycotoxins are not known to be produced. Ecology. This species (by either generic name) has rarely if ever been reported from the food literature. However, we have found Trichoconiella padwickii, reported as Alternaria padwickii, to be endemic in rice from Southeast Asia. It was present in 9 of 18 samples of Thai paddy rice, with up

5.28 Genus Trichoderma

to 60% of grains in individual samples showing infection, and was found in 12% of all individual grains examined (Pitt et al., 1994). The incidence in paddy rice from Indonesia (71% of samples, 19% of all particles) and the Philippines (72% of samples, 23% of all particles) was even higher. Individual rice samples from Indonesia had up to 69% of particles infected by T. padwickii, while incidence was up to 90% of grains in samples from the Philippines (Pitt et al., 1998a and our unpublished data). Ellis (1971) reported Trichoconiella padwickii (as A. padwickii) from rice grains from Egypt, India, Malaysia, Nigeria, Pakistan and Sabah, indicating that this species is very widespread. Occurrence on crops other than rice has rarely been reported. References. Ellis (1971); Jain (1975); Domsch et al. (1980).

5.28 Genus Trichoderma Pers. In this ubiquitous genus, reproduction is by small single-celled conidia produced from phialides which are arranged in irregular verticils, with the subterminal phialides borne more or less at right angles to the stipe. Colonies are low and spread rapidly. Mycelial growth is loose textured and characteristically develops irregularly, often appearing in tufts or isolated patches. Conidia, green in the common species, sometimes develop only after exposure to light. Trichoderma species have usually been considered to be soil fungi, but the genus is now also known to include plant symbionts and fungal pathogens. Some are also of increasing importance as human pathogens. Interest in the genus has increased sharply in recent years, as some species have been proposed as biocontrol agents against plant pathogenic fungi. Speciation in Trichoderma has proved to be exceptionally difficult. The most common species name in the literature, Trichoderma viride Pers., has often been incorrectly used (Rifai, 1969) and it is now considered to be a rare species (Jaklitsch et al., 2006). Conidia produced by T. viride have rough walls, whereas the majority of Trichoderma isolates produce conidia with smooth walls. Reports

139

in the literature on Trichoderma species should be treated with caution, as green-spored Trichoderma isolates have often been called T. viride regardless of the texture of their conidial walls. Because most Trichoderma isolates have similar morphology, taxonomy proved very difficult until the advent of molecular methods. The use of sequences from multiple gene loci has resulted in a large increase in species numbers, from 9 in 1970 to now over 80 (Samuels, 2006). Most species have been shown to have teleomorphs in Hypocrea, though few are found in laboratory culture (Samuels, 2006). The taxonomy of Trichoderma is still evolving: see Samuels (2006) for a clear overview of current understanding. The most commonly isolated Trichoderma species is Trichoderma harzianum Rifai (Bisset, 1991a; Samuels, 2006). This is the species we have encountered most frequently from foods, and it is probable that most foodborne Trichoderma isolates belong in T. harzianum. Care should be exercised in handling Trichoderma isolates in the laboratory. Conidia are small, are shed and dispersed readily, and contaminant colonies grow rapidly. Many Trichodermas produce powerful chitinases and cellulases and, in time, can overrun and destroy other cultures completely.

Trichoderma harzianum Rifai

Fig. 5.48

Colonies on CYA and MEA generally covering the whole Petri dish, often irregular in outline or with isolated tufts evident, of white to yellow mycelium, with bright to dull yellow green conidia developing over the whole surface or in patches or tufts; reverse pale or yellowish. Colonies on G25N less than 5 mm diam, with growth weak. At 58C, usually no growth, occasionally germination. No growth at 378C. Conidiophores consisting of highly branched structures, with a stipe bearing branches and the branches rebranching, all approximately at right angles, to form a pyramidal shape, with each branch bearing phialides irregularly; phialides ampulliform, commonly 5–7  3.0–3.5 mm, larger when borne apically, bearing conidia singly, not in chains; conidia often adhering in small clusters, spheroidal, subspheroidal, or sometimes broadly ellipsoidal,

140

5 Primary Keys and Miscellaneous Fungi

Fig. 5.48 Trichoderma harzianum (a) colonies on MEA, 7 d, 258C; (b) conidiophores, bar = 10 mm; ( c) conidia, bar = 5 mm

2.5–3.2(–4.0) mm diam or in length, smooth walled. Solitary aleurioconidia also formed by some isolates, spherical to broadly ellipsoidal, 6–8 mm diam. Distinctive features. Trichoderma harzianum produces conidiophores which are compactly branched in a pyramidal shape, the base being highly branched, and the apex usually bearing a solitary phialide. Conidia are smooth-walled, nearly spherical and less than 3.5 mm long. Taxonomy. The teleomorph of Trichoderma harzianum is Hypocrea lixii. This is not seen in laboratory culture (Samuels, 2006). Physiology. Because of the confusion in the literature over the identity of Trichoderma isolates, it is possible that much of the information reported for T. viride actually is based on studies on T. harzianum. Domsch et al. (1980) reported the optimum growth temperature for T. harzianum as approximately 308C, with a maximum near 368C. Our observations indicate a minimum growth temperature at or slightly above 58C. The minimum aw for growth is 0.91 at 258C (Griffin, 1963). Mycotoxins. Mycotoxin production by Trichoderma harzianum has not been reported. Ecology. Trichoderma harzianum is frequently isolated from cultivated and forest soils in all

parts of the world (Domsch et al., 1980). It has been reported from rotting tubers of cassava (Ekundayo and Daniel, 1973) and as a cause of spoilage in apples (Penrose et al., 1984). It has been reported from salmon and peas (data sheets, International Mycological Institute, Egham, Surrey). We found T. harzianum, at levels up to 1% of all grains or nuts examined, in maize from Thailand and Indonesia and sorghum from Indonesia. It was encountered at low levels in Philippine maize, Thai peanuts and Indonesian cowpeas (Pitt et al., 1993, 1994, 1998a). T. viride was not encountered at all. Trichoderma harzianum is no doubt widely distributed in foods, but it has often been reported as T. viride. References. Bisset (1984, 1991a, b; Samuels, 2006).

5.29 Genus Trichothecium Link Trichothecium is a distinctive genus with a single common species, T. roseum, characterised by sparse, pinkish colonies and conidia formed in a unique V-formation on long stipes. T. roseum is a

5.29 Genus Trichothecium

141

Fig. 5.49 Trichothecium roseum (a) colonies on CYA and MEA, 7 d, 258C; (b) conidiophores and conidia in situ, bar = 25 mm; ( c) conidia, bar = 10 mm

common saprophyte in damp and decaying habitats and is sometimes a weak pathogen of both plants and animals. Trichothecium roseum (Pers.) Link

Fig. 5.49

Colonies on CYA and MEA 50–60 mm diam, low and often sparse, characteristically coloured orange pink near salmon; reverse similarly coloured, or less intense, or brownish. Colonies on G25N barely macroscopic, 1–2 mm diam at most. At 58C, no germination to germination. No growth at 378C.

Conidiophores long, simple hyphae, bearing conidia at the tip successively, each formed as a blown-out cell below the previous one, offset from the hyphal axis and adhering loosely to form characteristic short, V-shaped chains; conidia approximately ellipsoidal to pyriform, with a single transverse septum, 16–20  8–12 mm, and with thin, smooth walls. Distinctive features. See genus preamble. Physiology. Domsch et al. (1980) reported growth temperatures for Trichothecium roseum as minimum 158C, optimum 258C and maximum 358C. Our data on growth at low temperature is at

142

variance with this, indicating a minimum growth temperature of 58C or near. Growth occurs down to 0.90 aw (Snow, 1949). Mycotoxins. Trichothecium roseum has been reported to produce trichothecene toxins, including trichothecin, trichothecolon and 12,13-epoxy-4(1-oxobut-2-enyloxy)-trichothec-9-ene (Konishi et al., 2003). Some of these toxins have been found in grapes and wheat in measurable quantities, but as spoilage or heavy growth of T. roseum is rare, they are unlikely to be significant in human health. However, their presence at quite low levels in grape musts has been reported to inhibit wine yeasts (Flesch et al., 1986). Ecology. As a ubiquitous and readily recognised saprophyte, Trichothecium roseum has been isolated from a variety of foods. Cereals are a common source, including barley, wheat, maize, sorghum and paddy rice (see Pitt and Hocking, 1997). Other important sources include apples (Valletrisco and Niola, 1983) and grapes (see Pitt and Hocking, 1997; Serra et al., 2005; Blancard et al., 2006). This species has caused spoilage of a wide variety of other fruits and vegetables from time to time, but is usually only a minor pathogen (Snowdon, 1990, 1991). It has also been isolated from meat products, cheese, beans, hazelnuts, pecans and pistachios (see Pitt and Hocking, 1997). We isolated T. roseum from sorghum, cashews, paddy and milled rice, peanuts, maize and soybeans in Southeast Asia, but always at a low incidence (Pitt et al., 1998a). In our experience, levels of T. roseum in foods other than fruits are usually low, and spoilage exceptional. Reference. Domsch et al. (1980).

5.30 Genus Ulocladium Preuss Ulocladium, in common with Alternaria and Stemphylium, produces conidia with both longitudinal and transverse septa. Those of Ulocladium are narrower at the base than the apex, and can be pointed at one end, to give a pyriform or apiculate shape, while those of Stemphylium have rounded ends and are usually symmetrical longitudinally. Conidia of Alternaria are definitely club-shaped (Simmons, 1967). Although this distinction has

5 Primary Keys and Miscellaneous Fungi

been maintained by Simmons (2007), molecular analyses have indicated that Ulocladium cannot be separated from Alternaria (Pryor and Gilbertson, 2000; de Hoog and Horre´, 2002; Pryor and Bigelow, 2003). Ulocladium botrytis is treated here as representative of the genus.

Ulocladium botrytis Preuss

Fig. 5.50

Colonies on CYA and MEA 60 mm or more diam, plane, dense to somewhat floccose, dark olive to very dark grey, reverse blue black. On G25N, colonies 6–10 mm diam, of dense olive to dark grey mycelium, reverse blue black. At 58C germination to formation of microcolonies. At 378C, colonies 15–20 mm diam, olive grey, reverse grey brown or blue black. On DCMA, colonies 30–40 mm diam, similar to those on CYA. Conidia borne singly from nodes or pores on knobby or geniculate dark conidiophores, on CYA and DCMA variable in size and shape, often roughly spherical, 9–12 mm diam, occasionally single celled, usually with four or more septa, but also characteristically ellipsoidal, 15–25  9–10 mm, and with lateral and longitudinal or irregular septa, usually with only a small basal protrusion but occasionally stalked and resembling Alternaria, dark brown, with thick, often very rough walls. Distinctive features. As noted above, the genus Ulocladium is distinguished from Alternaria and Stemphylium by conidial shape. Conidia of Ulocladium are obovoid, i.e. narrower at the base than at the apex. U. botrytis is the only species at all common in foods. Physiology. No physiological studies on Ulocladium species are known to us. Mycotoxins. No mycotoxins are known to be produced. None of 52 Ulocladium isolates produced altertoxins or tenuazonic acid, toxic metabolites produced by many Alternaria species (Andersen and Hollensted, 2008). Ecology. Ulocladium species are infrequently found in foods. They have been isolated from hazelnuts and walnuts, pistachios, peanuts and barley

5.30 Genus Ulocladium

143

Fig. 5.50 Ulocladium botrytis (a) colonies on CYA and MEA, 7 d, 258C; (b) conidiophores and conidia in situ, bar = 50 mm; (c) conidia, bar = 10 mm

(see Pitt and Hocking, 1997). U. chartarum can cause rots in melons (Snowdon, 1990). We isolated Ulocladium species, at low levels, from cashews, red

beans, maize, peanuts and soybeans (Pitt et al., 1993, 1994, 1998a). References. Simmons (1967).

“This page left intentionally blank.”

Chapter 6

Zygomycetes

Zygomycetes (currently the phylum Zygomycota) are relatively primitive fungi characterised by the production of solitary spores, zygospores, as their teleomorph. Zygomycetes of significance here are characterised by hyphae with few if any cross walls (septa): the hyphae are essentially unobstructed tubes. Absence of septa facilitates rapid translocation of nutrients and organelles such as mitochondria and nuclei between sites of growth, nutrient adsorption and spore formation. In consequence, Zygomycetes are also characterised by rapidity of growth. Many species are able to fill a Petri dish with loosely packed mycelium and to produce mature spores within 2 days of inoculation. Zygospores are large (usually greater than 30 mm diam), dark walled, distinctive bodies (Fig. 6.1) on which Zygomycetes rely for long-term survival. Formation of zygospores by the majority of species encountered in food spoilage requires mating by two strains, so zygospores are not commonly observed in the pure cultures used for identification. A few species such as Rhizopus sexualis do produce zygospores in pure culture, and this is a valuable taxonomic aid. Perhaps because they are physically difficult to separate from other more abundant spore types, little is known about the physiological properties of zygospores, such as resistance to heat and chemicals. Zygomycetes embrace a wide variety of fungi, with diverse habitats. Almost all require high water activities for growth. In damp situations, their rapid growth habit provides a selective advantage over most, more advanced fungi with septate hyphae. Many are found on dung or as insect pathogens. For a guide to genera of Zygomycetes which

can be grown in the laboratory, see O’Donnell (1979). The genera of Zygomycetes commonly found in foods are confined to a single order, Mucorales. Order Mucorales. The order Mucorales is monophyletic (having a single evolutionary origin) with all genera closely related genetically (White et al., 2006). However, classification within the order is less certain, as molecular studies have indicated poor correlation with traditional morphological taxonomy (O’Donnell et al., 2001). All of the families and many of the genera within the order cannot be sustained using molecular data. Family Mucoraceae. At the same time, O’Donnell et al. (2001) consider that all of the genera included here fit within modern concepts of the family Mucoraceae, which is now being broadened to include many of the other previously recognised families (White et al., 2006). As the task of reordering the genera within this family remains incomplete, the generic taxonomy provided here is based on traditional systems. Comments on the taxonomic status of each genus is included under that name as appropriate. Although some species within this family produce zygospores as a sexual state in culture, traditional classification has been based on asexual (anamorphic) reproduction. Asexual reproduction in the order Mucorales is primarily by sporangiospores, which are typically borne within sporangia. Sporangia (Fig. 6.2a) are closed sacs, borne on stipes (stalks). Stipes are often termed sporangiophores, although this name is more appropriately applied to the whole fruiting structure in line with conventional terminology in other asexual fungi.

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_6, Ó Springer ScienceþBusiness Media, LLC 2009

145

146

Fig. 6.1 Zygospore of Rhizopus sexualis, bar ¼ 50 mm

Stipes may be borne singly or in clusters from fertile hyphae, may be branched or unbranched or may grow out from beneath a terminal sporangium to produce a succession of sporangia (sympodial branching, Fig. 6.2b). Sporangia appear under the low-power microscope as small (less than l mm diam) brown to black spheres distributed on long stipes throughout the aerial mycelium. Sporangial walls disintegrate in age, releasing the sporangiospores. In microscopic mounts from 7 day old cultures, intact sporangia are rare: the structures mostly seen at stipe apices are columellae (Fig. 6.2c), formed within the sporangia and remaining intact in age. The manner in which columellae collapse after sporangial disintegration provides useful taxonomic information.

6 Zygomycetes

In one genus, Syncephalastrum, sporangiospores are borne in cylindrical merosporangia in a radial array on the columella surface (Fig. 6.2d). Under low magnifications, these structures have some resemblance to the fruiting structures of Aspergillus. In another, Thamnidium, sporangiospores are borne both in normal sporangia and in small sporangioles, formed in clusters (Fig. 6.2e), often on the same stipe as a sporangium. In Cunninghamella, sporangioles are also borne in clusters, on spicules, but never produce sporangiospores. Here the sporangiole itself acts as the reproductive unit. Some species in the Mucoraceae also produce chlamydoconidia as a second type of anamorph. These are cylindrical to spherical cells with relatively thick walls formed in hyphae and stipes, sometimes in great numbers (Fig. 6.2f). They probably function as a resting stage, more resistant to light, heat and desiccation than sporangiospores, and are probably analogous to similar structures formed by Hyphomycetes. Identifying genera in Mucoraceae. One theme of this book is to provide a standardised system for identification of food spoilage fungi. Where possible, identification is achieved after 7 days incubation, by a single macroscopic and microscopic inspection of a standard set of Petri dishes. The

Fig. 6.2 Anamorphic reproductive structures in genera of Mucorales: (a) sporangium; (b) sympodial branching; (c) columellae; (d) merosporangium; (e) sporangioles; (f) chlamydoconidia

6

Zygomycetes

keys to Mucoraceae hereunder are accordingly based on this system. To the experienced mycologist it will be obvious that because they grow so rapidly, isolates of Mucoraceae can be identified much earlier than 7 days. However, a second theme of this book is the facilitation of fungal identifications by the bacteriologist who does not instinctively recognise a Mucor from a Monascus, so the standard 7 day schedule has been maintained here. There is an unexpected bonus: the characteristic shapes of columellae collapsing in age simplifies recognition of some genera (Fig. 6.3). Significant genera. Seven genera in the family Mucoraceae are treated here: Absidia Tiegh., Cunninghamella Matr., Mucor P. Micheli: Fr., Rhizomucor (Lucet and Costantin) Vuill., Rhizopus Ehrenb., Syncephalastrum Schroter and Thamnidium ¨ Link. These genera are differentiated primarily by the morphology of their sporangiophores and sporangia. Rhizoids, mucous and contamination. Other useful features for distinguishing some of these genera are the production of rhizoids and the

147

secretion of mucous material. Rhizoids are short root-like structures produced by Rhizopus at the base of each sporangiophore (Fig. 6.3c). Mucor species do not produce rhizoids, whereas Absidia and Rhizomucor species produce them irregularly. As the name implies, Mucor produces sporangiospores in a layer of mucous which causes them to adhere to the colony when disturbed or picked with a needle. Absidia and Rhizomucor are similar, but Rhizopus produces dry spores. If a Mucor culture fills the Petri dish the lid can be removed with minimal disturbance to the culture; however, the rhizoids of Rhizopus adhere to the lid and part of the culture will detach with it. Beware! Carry out this operation away from your inoculating area. Dry spores, which are aerially dispersed when plates are opened, and very rapid growth make Rhizopus species a serious source of laboratory contamination. The seven genera of Mucoraceae considered here are keyed below and then treated in alphabetical order.

Fig. 6.3 Collapse of columellae in age: (a) little collapse (Mucor, Rhizomucor); (b) funnel shapes (Absidia); (c) umbrella shapes (Rhizopus)

148

6 Zygomycetes

Key to genera of the family Mucoraceae included here 1

All reproductive cells borne on spicules around vesicles Some or all reproductive cells sporangiospores, borne within sporangia (Fig. 6.2a, b, c)

2 3

2 (1)

Reproductive cells sporangiospores, borne in cylindrical sacs (merosporangia; Fig. 6.2d) (at low magnification reminiscent of Aspergillus) Reproductive cells single-celled spherical sporangioles

Syncephalastrum Cunninghamella

3 (1)

As well as sporangia, clusters of smaller sacs (sporangioles, Fig. 6.2e) present Only sporangia, or columellae derived from sporangia, present

4 (3)

Columellae retaining approximately spherical shape after sporangiospore discharge (Fig. 6.3a), sporangiospore walls smooth or spiny Columellae collapsing to form funnel or umbrella shapes, sporangiospore walls smooth or striate

5 (4)

Sporangiospores rarely exceeding 5 mm in long axis Sporangiospores commonly exceeding 5 mm in long axis

6 (4)

Columellae collapsing inwardly from the apex to form a funnel shape (Fig. 6.3b), sporangiospore walls smooth Columellae collapsing outwardly to form an umbrella shape (Fig. 6.3c), sporangiospore walls striate

6.1 Genus Absidia Tiegh. The genus Absidia produces a distinctive type of columella which widens gradually at the junction with the stipe, outside the circumference of the sporangium (Fig. 6.4b). In other Zygomycete genera considered here, the junction of stipe and

Thamnidium 4

5 6 Rhizomucor Mucor

Absidia Rhizopus

columella is abrupt, and the columella is wholly within the sporangial wall. In age, columellae of Absidia frequently collapse inward from the apex to form funnel-shaped structures. Absidia species form rhizoids, irregular root-like outgrowths at the bases of the stipes, but these are less conspicuous and less regular than in Rhizopus

Fig. 6.4 Absidia corymbifera (a) colonies on CYA, 7 days, 258C; (b) sporangia and columellae, bar ¼ 25 mm; (c) sporangiospores, bar ¼ 10 mm

6.2

Genus Cunninghamella

149

species. Only one Absidia species, Absidia corymbifera, is at all common in foods. Several recent authors have concluded that Absidia is polyphyletic (evolutionarily diverse) based on molecular studies (e.g. Voigt et al., 1999; O’Donnell et al., 2001; Kwasna et al., 2006) and that A. corymbifera is not closely related to most other species in the genus and would be better classified in Rhizomucor. However, this name remains in common use.

Absidia corymbifera (Cohn) Sacc. & Trotter

Fig. 6.4

recorded as a pathogen of peaches (Singh and Prashar, 1988). A. corymbifera is probably widespread in the tropics: Oyeniran (1980) recorded isolations from cocoa, palm kernels and maize. We have found it, always at low levels, in sorghum and mung beans from Thailand (Pitt et al., 1993, 1994); in peanuts, kemiri nuts, milled rice and coriander from Indonesia (Pitt et al., 1998a); and peanuts, paddy, milled rice, soybeans and black pepper from the Philippines (our unpublished data). References. Ellis and Hesseltine (1966); Nottebrock et al. (1974); de Hoog et al. (2000).

Absidia ramosa J.J. Ellis & Hesselt.

On CYA, colonies covering the Petri dish, low and sparse, white to pale brown or grey; reverse colourless. On MEA, colonies filling the whole Petri dish with deep floccose mycelium, coloured mid-grey by sporangia; reverse pale. On G25N, colonies 10–30 mm diam, sparse and floccose, coloured as on MEA. No growth at 58C. At 378C, colonies covering the whole Petri dish, similar to those at 258C. Sporangiophores borne from aerial hyphae, stipes sometimes irregularly branched; sporangia hyaline, 15–50 mm diam, appearing pyriform due to external conical columellae; columellae pyriform 10–30 mm diam, sometimes with small projections on the apices or with collarettes above the base, in age often collapsing inward from the apex to form funnel-shaped structures; sporangiospores hyaline, broadly ellipsoidal to spheroidal, 3–6 mm long, smooth walled. Distinctive features. See genus description. Physiology. Evans (1971) recorded growth temperatures for Absidia corymbifera as minimum 148C, maximum 508C and optimum near 408C. This species is able to germinate and grow down to 0.88 aw (Hocking and Miscamble, 1995). Growth in nitrogen (<1% O2) was similar to that in air (Hocking, 1990). Mycotoxins. Mycotoxin production has not been reported. Ecology. Absidia corymbifera is a weak human and animal pathogen, with a very wide host range and capable of infecting many body organs (de Hoog et al., 2000). From foods, most isolations have been from wheat, barley, malted barley and cereal products such as flour and bran. It has also been found in meat products and biltong, cassava, hazelnuts and sunflower seeds and, as A. ramosa, in pecans (see Pitt and Hocking, 1997). It has been

6.2 Genus Cunninghamella Matr. This genus is unusual in that sporangiophores give rise to sporangioles (small sporangia) which do not differentiate into sporangiospores but themselves act as the conidial stage. The sporangioles are borne on spicules (spikes) from vesicles; the vesicles are borne terminally or irregularly on the sporangiophores. A recent monograph by Zheng and Chen (2001) lists 15 species, and a molecular study of 12 of these showed good agreement with traditional speciation (Liu et al., 2001). Zheng and Chen (2001) report that a great deal of literature confusion exists over the names of species, in particular with the common species Cunninghamella bertholletiae and C. elegans. These two species are distinguishable by small morphological differences and by the fact that C. bertholletiae grows to 428C or more, while the maximum growth temperature for C. elegans is usually 358C. Although the name C. elegans is most commonly seen in the food literature, our experience indicates that C. bertholletiae is the most common species, along with C. echinulata. C. bertholletiae is described below: C. echinulata differs by the production of sporangioles with conspicuously spiny walls (Zheng and Chen, 2001).

Cunninghamella bertholletiae Stadel

Fig. 6.5

Colonies on CYA and MEA covering or filling the whole Petri dish, sparse, mycelium off-white to beige, reverse colourless to pale yellow. On G25N, ranging from no growth to colonies 15 mm diam, of sparse, translucent mycelium. At 58C, no

150

6 Zygomycetes

Fig. 6.5 Cunninghamella bertholletiae (a) colonies on CYA and MEA, 7 days, 258C; (b) developing sporangioles in situ, bar ¼ 50 mm; (c, d) vesicles and mature sporangioles, bar ¼ 10 mm

germination. At 378C, colonies similar to those on CYA at 258C, mycelium off-white to grey, reverse pale yellow to brown. Sporangiophores long, bearing solitary vesicles terminally and in irregular verticils subterminally or along the length; sporangioles borne from spicules, breaking at maturity, spherical to ellipsoidal, 7–11 mm diam or 9–13 mm long, with smooth to finely roughened walls. Zygospores not commonly seen in pure culture, fide Zheng and Chen (2001) irregularly ellipsoidal to spherical, brownish, 20–45 mm diam, with rough walls. Distinctive features. See genus description. Taxonomy. Most Cunninghamella species are heterothallic, producing zygospores only when two

mating types from the same species are grown together. Physiology. Cunninghamella bertholletiae grows up to 42–458C. Growth in soil is confined to above – 90 bars (ca. 0.93 aw) (Kouyeas, 1964). Mycotoxins. Mycotoxins are not produced. Ecology. In common with other Cunninghamella species, C. bertholletiae mostly occurs in tropical and subtropical soils (Zheng and Chen, 2001). C. echinulata was a cause of rotting in kola nuts (Adebajo, 1994). We have seen C. bertholletiae at low levels in maize from the Philippines and peanuts from Indonesia (Pitt et al., 1998a and unpublished). References. O’Donnell (1979); Domsch et al. (1980).

6.3

Genus Mucor

151

6.3 Genus Mucor P. Micheli: Fr. As traditionally described, Mucor is a very common and widespread genus in nature, occurring in soils, decaying vegetation, dung and many other moist habitats where rapidly growing fungi have an advantage. Recent molecular studies (Voigt et al., 1999; O’Donnell et al., 2001; Kwasna et al., 2006) have indicated that Mucor includes at least two phylogenetically distinct groups. However, the species included here, insofar as they have been examined, appear to fall into the major Mucor clade, so the traditional description is used here. Unlike those of Absidia, sporangia of Mucor have columellae borne wholly within the sporangial wall; the columellae collapse irregularly, if at all, in age (Fig. 6.3a). Unlike Rhizopus species, Mucors do not produce rhizoids. In species of interest here, sporangiospores are longer than 5 mm and have walls

which are smooth or spiny, but not striate. This is in contrast to Rhizopus species, which commonly produce sporangiospores with striate walls. Some Mucor species are able to grow and weakly ferment under anaerobic conditions and occasionally cause spoilage of beverages in this manner. Growth is yeast-like in appearance (Fig. 6.6), although individual cells are much too large to be mistaken for a true yeast. Inoculation of such cells onto aerobic media produces normal growth. Yeast-like growth has also been reported to occur when Mucor and some related genera grow in the presence of high sodium chloride concentrations (Tresner and Hayes, 1971). At least 20 species of Mucor have been reported from foods. Five appear to be most significant: Mucor circinelloides, M. hiemalis, M. piriformis, M. plumbeus and M. racemosus. These species are keyed and described below. General reference. Schipper (1978a).

Key to Mucor species included here 1

Strong growth at 378C No growth at 378C

2 (1)

Columellae frequently with small irregular projections on the apices; sporangiospores with minute spines Columellae without apical projections; sporangiospores smooth walled

M. plumbeus 3

3 (2)

Columellae 50–100 mm diam; growth on G25N weak or absent Columellae usually less than 50 mm diam; colonies on G25N greater than 10 mm diam

M. piriformis 4

4 (3)

Chlamydoconidia abundant, often dominating microscopic appearance Chlamydoconidia present in low numbers or absent

M. racemosus M. hiemalis

Mucor circinelloides Tiegh.

M. circinelloides 2

Fig. 6.7

On CYA, colonies 60 mm diam or more, often spreading across the whole Petri dish, but growth

Fig. 6.6 M. plumbeus showing yeast-like growth, in liquid culture under anaerobic conditions, bar ¼ 25 mm

relatively low and sparse, appearing pale grey or yellowish; reverse uncoloured. On MEA, colonies filling the whole Petri dish, in colours similar to those on CYA. On G25N, colonies 15–25 mm diam, low and relatively dense, golden yellow in both obverse and reverse. At 58C, colonies 4–10 mm diam, low and sparse. At 378C, colonies 20–40 mm diam, sparse and floccose, with colours more brown than at 258C. Sporangiophores borne from aerial hyphae, stipes commonly branched, often sympodially, sporangia spherical, 25–50 mm diam, sometimes up to 80 mm; columellae roughly spherical, up to 50 mm diam; sporangiospores hyaline, ellipsoidal, mostly 4.5–7 mm long, smooth walled. Chlamydoconidia uncommon, spherical, cylindrical or rather irregular, up to 15 mm diam. Zygospores not formed in pure culture.

152

6 Zygomycetes

Fig. 6.7 Mucor circinelloides (a) colonies on CYA, 7 days, 258C; (b) columellae; (c) sporangia; (d) chlamydoconidia; (e) sporangiospores; (f) sporangium; (g) sporangiospores, all bars ¼ 10 mm

Distinctive features. Mucor circinelloides has numerous characters in common with M. hiemalis. Unlike M. hiemalis, M. circinelloides grows well at 378C (20 mm or more in 7 days), grows weakly at 58C (less than 10 mm vs. greater than 20 mm in 7 days) and commonly produces sympodially branched stipes. Taxonomy. DNA sequence data from the ITS1 and 2 regions indicate that Mucor circinelloides is correctly placed in the genus Mucor (Kwasna et al., 2006). Physiology. Tresner and Hayes (1971) reported that Mucor circinelloides grew in media containing 15% (w/v) NaCl (=0.90 aw) but not 20% (=0.86 aw). On a medium with glucose as controlling solute, this species germinated and grew down to 0.90 aw (Hocking and Miscamble, 1995). Mycotoxins. This species is not known to produce mycotoxins.

Ecology. This species has been reported frequently as an animal pathogen and occasionally as a human pathogen (de Hoog et al., 2000). It can cause spoilage in cheese and yams (see Pitt and Hocking, 1997) and can be pathogenic on mangoes (Johnson et al., 1990) and peaches (Restuccia et al., 2006). It has been isolated from meat, hazelnuts and walnuts and also from maize, mung beans, soybeans and barley (Pitt et al., 1993, 1994, 1998a; see Pitt and Hocking, 1997). References. Schipper (1976); Domsch et al. (1980).

Mucor hiemalis Wehmer

Fig. 6.8

On CYA, colonies spreading across, and sometimes filling, the whole Petri dish, growth relatively

6.3

Genus Mucor

153

Fig. 6.8 Mucor hiemalis (a) colonies on CYA, 7 days, 258C; (b) sporangia and columellae, bar ¼ 25 mm; (c) sporangiospores, bar ¼ 10 mm

sparse, greyish; reverse pale. On MEA, colonies filling the whole Petri dish, relatively dense, greyish to distinctly yellow; reverse yellow to golden yellow. On G25N, colonies 10–15 mm diam, moderately dense, bright yellow in both obverse and reverse. At 58C, colonies 20–30 mm diam, low and sparse. No growth at 378C. Sporangiophores borne aerially, stipes generally unbranched, less commonly sympodially branched; sporangia up to 60 mm diam; columellae ellipsoidal, 15–30 mm diam; sporangiospores hyaline, narrowly to broadly ellipsoidal or reniform (kidney shaped), 5–11 mm long, smooth walled. Chlamydoconidia uncommon, spherical to cylindrical or irregular, up to 15 mm diam. Zygospores not formed in pure culture. Distinctive features. Mucor hiemalis is similar to M. circinelloides, but grows more rapidly at 58C and does not grow at 378C. Moreover, it produces larger sporangiospores which are sometimes reniform and usually has unbranched stipes. Taxonomy. DNA sequence data from the ITS1 and 2 regions indicate that Mucor hiemalis is correctly placed in the genus Mucor (Kwasna et al., 2006). Physiology. Growth of some Mucor hiemalis isolates occurs at temperatures below 08C (Joffe, 1962). Mycotoxins. Mycotoxin production has not been reported.

Ecology. Mucor hiemalis has been reported as a rare cause of cutaneous mycosis (de Hoog et al., 2000). This species causes rots in guavas (Ito et al., 1979), carrots and cassava (Snowdon, 1991). It has been reported from spoilage of yoghurt due to inward collapse of containers (Foschino et al., 1993) and cheese (Hayaloglu and Kirbag, 2007). It has also been reported from fresh vegetables (Lugauskas et al., 2005), from chestnuts (Jermini et al., 2006), hazelnuts and soybeans (see Pitt and Hocking, 1997), from wheat based fast foods in Nigeria (Fapohunda and Ogundero, 1990), from preprepared airline food in Egypt (Saudi and Mansour, 1990) and from chocolate confectionery in Italy (Dragoni et al., 1989). We have isolated it at low levels from Thai and Indonesian maize and paddy rice (Pitt et al., 1993, 1998a). Reference. Schipper (1973).

Mucor piriformis A. Fisch.

Fig. 6.9

On CYA, colonies low and sparse, spreading across the Petri dish or discrete; mycelium colourless, overall colour buff from sporangia; reverse pale. On MEA, colonies 35–60 mm diam, coloured grey or brownish; reverse pale. On G25N, colonies less than 7 mm diam, or growth absent. At 58C, colonies

154

6 Zygomycetes

Fig. 6.9 Mucor piriformis (a) colonies on MEA, 7 days, 258C; (b) sporangia, bar ¼ 25 mm; (c) columellae, bar ¼ 25 mm; (d) sporangiospores, bar ¼ 10 mm; (e) developing sporangium, bar ¼ 25 mm; (f) sporangiospores, bar ¼ 10 mm

30–45 mm diam, low and sparse, or with some aerial hyphae. No growth at 378C. Sporangiophores on CYA borne from surface hyphae, stipes short, broad, often sympodially branched and with encrusted walls; sporangia up to 150 mm diam, with spinulose walls; columellae spheroidal to short cylindroidal, 25–80(–100) mm diam or in length, the larger ones collapsing irregularly; sporangiospores hyaline, spherical to broadly ellipsoidal, 6–12(–20) mm diam or in long axis, smooth walled. Chlamydoconidia uncommon; zygospores not formed in pure culture. Distinctive features. Mucor piriformis grows rapidly on CYA at 58C, but relatively poorly under the standard 258C incubation conditions. This is especially noticeable on G25N, where growth is weak or absent. Mycelium on CYA at

258C is often contorted and highly branched. Sporangia, columellae and many sporangiospores are larger than those of other common Mucor species. Physiology. Mucor piriformis is a psychrophile. Cardinal temperatures are minimum, near 08C, optimum 20–218C, maximum 268C (Michailides and Spotts, 1990). Mycelium and sporangiospores were inactivated by heating to 43–46 and 52–558C, respectively (Michailides and Ogawa, 1989). Mycotoxins. Mycotoxin production has not been reported. Ecology. Mucor piriformis is a destructive pathogen of fresh strawberries (Snowdon, 1990; Pitt and Hocking, 1997) and causes rotting of cold stored pears, apples and tomatoes (see Pitt and Hocking, 1997), plums (Borve and Vangdal, 2007) and yams (Amusa and Baiyewa, 1999; Iwata, 2006). A variety of

6.3

Genus Mucor

treatments for control of pear spoilage have been proposed: dips in hot water (Michailides and Ogawa, 1989), a hot water pressure process (Spotts et al., 2006), salt and surfactant solutions (Spotts and Cervantes, 1989), prestorage drench in thiabendazole (Lennox et al., 2004) or treatment of wash water with chlorine dioxide or peracetic acid (Roberts and Reymond, 1994; Mari et al., 2003). This species has recently been reported as a dominant species in the mycoflora of cassava and yam chips (Gnonlonfin et al., 2008). References. Schipper (1975); Michailides and Spotts (1990).

155

Mucor plumbeus Bonord.

Fig. 6.10

Mucor spinosus Tiegh.

On CYA and MEA, colonies at least 50 mm diam, low to deep, often spreading across the Petri dish; mycelium colourless, overall colour pale to deep grey from sporangia; reverse colourless. Colonies on G25N 20–35 mm diam, low, moderately dense, white to pale yellow brown; reverse pale. At 58C, colonies 8–15 mm diam, low and sparse. No growth at 378C. Sporangiophores borne from surface or aerial hyphae, stipes unbranched or branched sympodially,

Fig. 6.10 Mucor plumbeus (a) colonies on CYA, 7 days, 258C; (b) sporangia, bar ¼ 25 mm; (c) columellae, bar ¼ 25 mm; (d) chlamydoconidia, bar ¼ 25 mm; (e) sporangiospores, bar ¼ 10 mm; (f) sporangia, bar ¼ 25 mm; (g) sporangiospores, bar ¼ 10 mm

156

sporangia dark greyish brown, up to 80 mm diam, with spiny walls; columellae pyriform to ellipsoidal or short cylindroidal, up to 50  30 mm, often with irregular projections at the apices; sporangiospores brown, spheroidal, commonly 7–8(–12) mm diam, with walls rough or minutely spiny. Chlamydoconidia uncommon; zygospores not formed in pure culture. Distinctive features. Mucor plumbeus is distinguished by its grey colour, columellae with irregular apical projections and brown sporangiospores with rough or spiny walls. Taxonomy. DNA sequence data from the ITS1 and 2 regions indicate that Mucor plumbeus is correctly placed in the genus Mucor (Kwasna et al., 2006). Physiology. Panasenko (1967) reported growth of Mucor plumbeus from 4 or 5 to 358C, with an optimum of 20–258C. The minimum aw for growth was reported to be 0.93 by Snow (1949). Growth in N2 (<1% O2) was 80% of that in air; some growth occurred in an atmosphere of 97% CO2, with only trace amounts of O2 (Hocking, 1990). As measured by colony diameters, growth on cheddar cheese in an atmosphere of 20% CO2 and 5% O2 was 50% of that in air, and growth in 20% CO2 and 1% O2, 40% CO2 and 5% O2 and 40% CO2 and 1% O2 was 40, 50 and 30% of that in air, respectively (Taniwaki et al., 2001a). Mycotoxins. Mycotoxins are not known to be produced. Ecology. Mucor plumbeus has been reported to spoil cheese (Northolt et al., 1980; Devoyod, 1988)

6 Zygomycetes

and has been observed to cause anaerobic spoilage of apple juice in our laboratory. Meat (Gros et al., 2003; Pitt and Hocking, 1997), nuts and cereals are other commodities from which this species, and its synonym M. spinosus, have been reported (see Pitt and Hocking, 1997). It was isolated at low levels from black rice in Thailand, soybeans in the Philippines and coriander in Indonesia (Pitt et al., 1994, 1998a). Reference. Schipper (1976).

Mucor racemosus Fresen.

Fig. 6.11

On CYA and MEA, colonies spreading across the Petri dish, low to moderately deep; mycelium colourless, overall colour light to mid brown from sporangia and chlamydoconidia; reverse light brown. On G25N, colonies 25–40 mm diam, low, moderately dense, similar in colour to those on CYA. At 58C, colonies 12–25 mm diam, low and sparse. No growth at 378C. Sporangiophores borne from surface or aerial mycelium, stipes branched sympodially or irregularly, sporangia up to 80 mm diam, light brown, with encrusted walls; columellae ellipsoidal to pyriform, up to 40 mm long; sporangiospores hyaline to pale brown, broadly ellipsoidal to subspheroidal, commonly 5–8 mm diam, smooth walled. Chlamydoconidia and arthroconidia formed abundantly, 5–20 mm or more in diam or long axis. Zygospores not formed in pure culture.

Fig. 6.11 Mucor racemosus (a) colonies on CYA, 7 days, 258C; (b) columellae, bar ¼ 25 mm; (c) chlamydoconidia, bar ¼ 25 mm; (d) sporangiospores, bar ¼ 10 mm

6.4

Genus Rhizomucor

Distinctive features. Mucor racemosus is similar in many respects to M. plumbeus. M. racemosus differs by faster growth on G25N, brown not grey colony colouration, the much greater abundance of chlamydoconidia, smooth sporangiospores and the absence of irregular projections on columellae. Taxonomy. DNA sequence data from the ITS1 and 2 regions indicate that Mucor racemosus is correctly placed in the genus Mucor (Kwasna et al., 2006). Physiology. Mucor racemosus grows between –3 or –48C and 30–358C, with an optimum of 20–258C (Panasenko, 1967). The minimum aw for growth is 0.92 (Panasenko, 1967). Mycotoxins. No mycotoxins are produced. Ecology. Mucor racemosus is responsible for a spongy soft rot of cool stored sweet potatoes, potatoes and citrus (Chupp and Sherf, 1960). It has been reported, along with M. hiemalis, from spoilage of yoghurt due to inward collapse of containers (Foschino et al., 1993) and as a cause of spoilage of cheese (Devoyod, 1988; and in our laboratory) and cheesecake (Piskorska-Pliszczynska and BorkowskaOpacka, 1984). It has also been found contaminating cheese (Kivanc¸, 1992; Lund et al., 1995; Cantoni et al., 2003), salami (Cantoni et al., 2007), raisins (Youssef et al., 2000), frozen and processed meats, salted horse meat, fermenting cacao beans, maize, barley, soybeans and paddy rice (see Pitt and Hocking, 1997). Reference. Schipper (1976).

157

6.4 Genus Rhizomucor (Lucet and Costantin) Vuill. Schipper (1978b) revived the genus Rhizomucor for three species previously accepted in Mucor, but distinguished by the production of stolons and by their thermophilic nature. Molecular studies (Voigt et al., 1999) support this concept. Rhizomucor species are significant in foods mainly in tropical regions. The most common species in foods is Rhizomucor pusillus; R. miehei is also mentioned.

Rhizomucor pusillus (Lindt) Schipper Fig. 6.12 Mucor pusillus Lindt

On CYA, colonies sometimes 25–35 mm diam, of colourless to white floccose mycelium surmounted by brown sporangia, or sometimes covering the whole Petri dish and then similar to that on MEA. On MEA, colonies covering the whole Petri dish, low and relatively sparse, pale to mid-grey; reverse pale, yellowish or greenish grey. On G25N, growth sparse, not exceeding 5 mm diam, or absent. No growth at 58C. At 378C, colonies similar to those at 258C on MEA, but more dense, brown to dark grey. Sporangiophores borne from surface hyphae, stipes sometimes appearing unbranched, but usually extensively and irregularly branched; poorly formed rhizoids sometimes apparent but not adjacent to the

Fig. 6.12 Rhizomucor pusillus (a) colonies on CYA, 7 days, 258C; (b) columellae, bar ¼ 25 mm; (c) sporangia, bar ¼ 25 mm; (d) sporangiospores, bar ¼ 10 mm

158

6 Zygomycetes

stipe bases, a distinction from Rhizopus; sporangia spherical, brown or grey, 40–60(–80) mm diam; columellae spherical, ellipsoidal or pyriform, 20–45 mm diam, sometimes collapsing irregularly (similar to Mucor species); sporangiospores hyaline, spherical to broadly ellipsoidal, 3–4 mm diam, smooth walled. Zygospores produced by occasional isolates, black, broadly ellipsoidal, 60–70 mm diam. Distinctive features. Rhizomucor species are most readily distinguished by rapid growth at 378C and by their sporangiospores, which are small (less than 5 mm diam) and smooth walled. Physiology. Rhizomucor pusillus is a thermophile. Growth has been reported at temperatures as high as 608C (Crisan, 1973). Optimum growth conditions are 37–428C, with a lower limit of 208C (Panasenko, 1967; Evans, 1971). Mycotoxins. Mycotoxin production is unknown. Ecology. Rhizomucor pusillus is a frequent agent of bovine mycotic abortion and a rare cause of human zygomycosis (de Hoog et al., 2000). This species appears to be of widespread though uncommon occurrence in foods. Reported sources include cereals (Lugauskas et al., 2006), kola nuts (Adebajo and Popoola, 2003), various spices (Mandeel, 2005), olives (Roussos et al., 2006), meat products, pecans, hazelnuts and walnuts, sunflower seeds and various tropical products (see Pitt and Hocking, 1997). It is used in cheese manufacture in Japan (Koizumi, 2001). We isolated R. pusillus from mung beans in Thailand and kemiri nuts in Indonesia (Pitt et al., 1994, 1998a). Additional species. Rhizomucor miehei (Cooney and R. Emers.) Schipper (synonym M. miehei Cooney and R. Emers.) has occasionally been isolated from foods (Kuthubutheen, 1979; Ogundero, 1981). It is similar in most characters to R. pusillus, including thermophily. R. miehei produces sporangia up

to 50(–60) mm diam with spiny walls, with columellae rarely larger than 30 mm diam. Reference. Schipper (1978b).

6.5 Genus Rhizopus Ehrenb. The genus Rhizopus needs little introduction to food microbiologists, because R. stolonifer is among the most obvious moulds encountered on Petri dishes inoculated with food materials. Coarse, rampant growth and rapidly maturing spores make this species a source of endless contamination to the unwary laboratory worker. Other Rhizopus species are less common in foods and cause less nuisance. Rhizopus species, especially R. oligosporus, have been used for millennia to modify basic foods in the Orient. The subject of fermented foods is outside the scope of this book: for further information see, for example, Hesseltine (1965), Gray (1970), Beuchat (1987) or Rombouts and Nout (1995). Rhizopus is distinguished from other genera in the order Mucorales by the formation of rhizoids (short root-like appendages), which are conspicuous at the base of the sporangiophores; by columellae which often collapse into umbrella shapes in age; and by dry sporangiospores with striate walls. The most recent complete taxonomic treatment is by Zheng et al. (2007a) who accepted 10 species. Molecular phylogenetic analyses have recently been carried out by Abe et al. (2006) and Liou et al. (2007), with good agreement with the morphological conclusions by Zheng et al. (2007a). Five species are considered here and keyed below: R. microsporus, R. oligosporus, R. oryzae, R. sexualis and R. stolonifer. Because of its ubiquitous nature, R. stolonifer is described first.

Key to Rhizopus species included here 1

Abundant zygospores present on CYA and MEA Zygospores not produced

2 (1)

Rapid growth at 378C; spores commonly less than 8 mm diam Growth at 378C weak or absent; spores commonly 8–20 mm diam

3 (2)

Columellae up to 100 mm diam; spores 5–8 mm long Columellae not more than 80 mm diam; spores less than 5 mm long

4 (3)

Columellae up to 75 mm diam; spores less than 4 mm long, with spinose walls; source food fermentations and fermented foods Columellae less than 40 mm diam; spores 3–5 mm long, with finely striate walls; source other types of foods

R. sexualis 2 3 R. stolonifer R. oryzae 4

R. oligosporus R. microsporus

6.5

Genus Rhizopus

Rhizopus stolonifer (Ehrenb.: Fr.) Lindner

159

Fig. 6.13

Rhizopus nigricans Ehrenb.

On CYA, colonies covering the whole Petri dish, sometimes low and sparse, with black sporangia only at the margins, sometimes filling the whole Petri dish and then similar to those on MEA; reverse pale. On MEA, colonies filling the whole Petri dish with floccose white mycelium bearing conspicuous sporangia, at first white, then with maturation rapidly becoming black, either distributed uniformly or concentrated at dish peripheries; reverse uncoloured. On G25N, colonies similar to that on MEA but less dense. At 58C, spores barely germinating. At 378C, usually no growth, sometimes colonies up to 15 mm diam, very thin and sparse. Sporangiophores borne in groups of three to five from clusters of rhizoids, stipes unbranched, robust and up to 3 mm long, with brown walls; sporangia 100–350 mm diam, usually spherical; columellae roughly spherical, up to 200 mm diam, in age often collapsing downwards and outwards to produce umbrella shapes; sporangiospores commonly 8–20 mm in long axis, pale brownish, with striate walls. Distinctive features. Rhizopus stolonifer is distinguished by its habit, with coarse hyphae and rampant growth at 258C, and by sporangia which are white when first formed but which change to black

with maturity. Sporangiospores are large, with striate walls. In contrast to that at 258C, growth at 5 and 378C is weak or absent. Taxonomy. Twenty nine isolates identified as Rhizopus stolonifer or possibly related species were phylogenetically analysed by RAPD analysis (Va´gvolgyi ¨ et al., 2004) and ca. 30 different isolates by Liou et al. (2007) using DNA sequences from the D1/D2 of rDNA. These analyses divided R. stolonifer and some varieties into two or four clades. In each case, the isolates commonly seen in foods remained in R. stolonifer. Physiology. Rhizopus stolonifer has been reported to grow from 4.5 or 58C up to 308C (Schipper, 1984) or 35–378C (Pierson, 1966, and our observations), with an optimum near 258C. However, Zheng et al. (2007a), who tested 36 isolates of R. stolonifer, reported maximum growth temperatures of only 30–328C. This species germinated down to 0.84 aw at 258C, but growth was very slow, and was absent at 308C (Hocking and Miscamble, 1995). It produced the fastest mycelial growth we have recorded: at 258C and an aw in excess of 0.99, the radial growth rate reached 2 mm/h (i.e. nearly 5 cm/day). Like some other species in the Mucorales, R. stolonifer can grow under anaerobic conditions (Stotzky and Goos, 1965); however, an atmosphere of 80% O2 + 20% CO2 halved its growth rate at 258C after 40 h (Hoogerwerf et al., 2002).

Fig. 6.13 Rhizopus stolonifer (a) colonies on CYA, 7 days, 258C; (b) sporangiophores with rhizoids and collapsed columellae, bar ¼ 25 mm; (c) sporangiospores, bar ¼ 10 mm

160

Mycotoxins. Rhizopus stolonifer (a single isolate tested) was moderately toxic to ducklings (Rabie et al., 1985). The possible toxins involved have not been elucidated. Ecology. Rhizopus stolonifer is by far the most commonly occurring species of the order Mucorales in foods. It causes destructive rots of a wide range of fruits (Snowdon, 1990, 1991). The most important are fresh berries, especially strawberries (Beneke et al., 1954; Dennis et al., 1979; Harris and Dennis, 1980), and stone fruits of all kinds (Snowdon, 1990; Singh and Shukla, 2005). Tomatoes, eggplant and capsicums are also seriously affected in some countries (Kassim, 1987; Snowdon, 1991). These diseases occur postharvest and are commonly known as ‘‘transit rot’’. Whole cases of fruit can decay in just a few days. Spoilage is usually initiated through damage and then spreads by contact. Control usually relies on preharvest or postharvest sprays or dips of fungicides. Thiabendazole (Benlate) or 2,6-dichloro-4-nitroaniline (Dichloran or Botran) provides reasonable control (Snowdon, 1990, 1991 and see Pitt and Hocking, 1997). Other treatments suggested include hot water dips and irradiation (Nicoue et al., 2004), ultraviolet light (Pan et al., 2004) and chlorine dipping (Sabaa-Srur et al., 1993). Control of postharvest rot has been satisfactorily achieved with applications of yeasts such as Metschnikowia fructicola (Karabulut et al., 2004) and Cryptococcus laurentii (Zhang et al., 2007a). Transgenic tomatoes with reduced polygalacturonase activity appear to have a higher resistance to transit rot (Sanders et al., 1992). Pectic enzymes of Rhizopus stolonifer survive the canning process normally applied to fruit. If even a small proportion of infected fruit is processed, these enzymes can cause softening and spoilage of canned apricots (Harper et al., 1972). Many types of vegetables are also susceptible to spoilage by Rhizopus stolonifer (Snowdon, 1991). Beans and peas, carrots, sweet potatoes, yams and cassava (see Pitt and Hocking, 1997) are all highly susceptible. Isolation of this species has been reported from many other food sources, including wheat and barley (Lugauskas et al., 2006), soybeans (Tariq et al., 2005), peanuts (Gachomo et al., 2004), hazelnuts (Senser, 1979), pecans and bran (see Pitt and Hocking, 1997), spices (Mandeel,

6 Zygomycetes

2005), cheese (Hayaloglu et al., 2007) and meat products (see Pitt and Hocking, 1997). R. stolonifer is a cosmopolitan fungus, particularly in tropical and subtropical regions, in almost all kinds of fresh, moist or partially dried foods. References. Schipper (1984); Zheng et al. (2007a).

Rhizopus microsporus Tiegh.

Fig. 6.14

On CYA and MEA, colonies covering or filling the whole Petri dish, of fine, pale grey mycelium, becoming dark grey as sporangia mature; reverse pale to dull yellow. On G25N, germination to microcolony formation. No growth at 58C. At 378C, colonies filling the whole Petri dish, similar to those on CYA or MEA at 258C. Sporangiophores borne singly or in groups of two to three from clusters of poorly formed rhizoids, stipes unbranched, relatively short, commonly 200–500 mm long, with brown walls; sporangia black, 100–200 mm diam, usually spherical; columellae 20–35 mm diam, spherical, some remaining so in age, others collapsing downwards, to form umbrella shapes; sporangiospores ellipsoidal, 3–5 mm long, with thin, striate walls. Distinctive features. Rhizopus microsporus differs from R. stolonifer by shorter stipes and smaller columellae, by the absence of white immature sporangia and by strong growth at 378C. It is distinguished from R. oryzae by production of smaller columellae and smaller spores. R. oligosporus is very similar to R. microsporus, but produces larger columellae and smooth to spinose spores. Taxonomy. Sporangia up to 80 mm and spores up to 6.5(–7.5) mm long were described by Schipper and Stalpers (1984). This species was divided into six varieties by Zheng et al. (2007a), one of which, Rhizopus oligosporus, is considered here to be a separate species. Other varieties are uncommon or unknown from foods. Physiology. Rhizopus microsporus grows up to 46–488C (Schipper and Stalpers, 1984; Zheng et al., 2007a). The lower limit for growth is 0.90 aw (Hocking and Miscamble, 1995). Mycotoxins. Wilson et al. (1984) were the first to report the production of rhizonin A, a nonspecific hepatotoxin, by Rhizopus microsporus. However, a recent study demonstrated that the toxin was not

6.5

Genus Rhizopus

161

Fig. 6.14 Rhizopus microsporus (a) colonies on CYA, 7 days, 258C; (b) sporangiophores with rhizoids, bar ¼ 25 mm; (c) sporangiospores, bar ¼ 10 mm

actually synthesised by the fungus, but by a bacterial symbiont belonging to the genus Burkholderia (Partida-Martinez et al., 2007). This finding is of interest as species of Rhizopus are frequently used in the food industry for the preparation of fermented foods such as tempe and sufu. Ecology. Rhizopus microsporus causes human zygomycoses, particularly cutaneous and gastrointestinal infections (de Hoog et al., 2000). This species appears to be uncommon in foods from temperature zones; however, we isolated it quite frequently from tropical commodities. It was found in 18% of maize samples from the Philippines and from 3% of all kernels examined. It was also present, at lower numbers, in peanuts, paddy rice, soybeans and black pepper from the Philippines, and maize, peanuts, paddy and milled rice, soybeans, mung beans and coriander from Indonesia (Pitt, 1998a and our unpublished data). Reference. Schipper and Stalpers (1984); Zheng et al, (2007a).

Rhizopus oligosporus Saito

Fig. 6.15

Colonies on CYA 50–60 mm diam, not usually covering the whole Petri dish, low, plane and sparse, with ill-defined margins, pale brown, sporangia sparsely produced, brown, sometimes enclosed in clear to brown droplets; reverse uncoloured to pale brown. Colonies on MEA covering the whole Petri dish, sometimes reaching, and adhering to, the lid, coloured dark grey to black, sporangia abundant, black; reverse uncoloured. On G25N, sometimes germination. At 58C, no germination. At 378C, colonies covering the whole Petri dish in a low, very sparse, often cobwebby growth; reverse uncoloured. Sporangiophores best observed on MEA, stipes 150–400 mm long, usually with well developed short rhizoids at or near the base, terminating in dark sporangia, 80–120 mm diam, at 7 days usually broken with spores dispersed; columellae persistent, often split, but remaining spheroidal to pyriform,

162

6 Zygomycetes

Fig. 6.15 Rhizopus oligosporus (a) colonies on MEA, 7 days, 258C; (b) sporangiospores, bar ¼ 10 mm; (c) developing sporangia, bar ¼ 25 mm; (d) sporangia, bar ¼ 25 mm

25–75 mm diam or long; sporangiospores spherical to subspheroidal, 3.0–3.5 mm diam, with thin, finely spinose walls. Distinctive features. In common with Rhizopus microsporus, R. oligosporus differs from R. stolonifer by shorter stipes and smaller columellae, by the absence of white immature sporangia and by strong growth at 378C. It is distinguished from R. oryzae by production of smaller columellae and smaller spores. R. oligosporus produces larger columellae than isolates of R. microsporus and forms smooth to spinose spores. Taxonomy. Schipper and Stalpers (1984) reduced Rhizopus oligosporus to the status of a variety of R. microsporus, and this was maintained by Zheng et al. (2007a). However, this taxon deserves species status as a domesticated species used in, and

probably confined to, food fermentations. Its derivation from R. microsporus seems likely, as the two taxa share many properties. Using a detailed light and scanning electron microscopy study, Jennessen et al. (2008) have illustrated criteria for separating R. oligosporus from related species. Sporangiospores of R. oligosporus are often larger and less regularly shaped than those of other species. This is likely to be the result of domestication (Jennessen et al., 2008). Physiology. Maximum growth temperature of 13 isolates was 45–498C (Zheng et al., 2007a). Mycotoxins. No mycotoxins are known to be produced. Weak toxicity to ducklings was reported by Rabie et al. (1985). However, only a single isolate was tested. Ecology. Rhizopus oligosporus is used in food fermentations, the most notable product being tempeh,

6.5

Genus Rhizopus

which is produced in Southeast Asian countries, especially in Indonesia. Cooked, dehulled soybeans are soaked for 2–3 days, then inoculated from a previous batch or, more commonly now, with a starter culture, usually R. oligosporus. Fermentation takes place under ambient (tropical) conditions, preferably with air temperatures of 25–288C for 36–48 h (Hesseltine, 1965, 1991; Ko and Hesseltine, 1979; Beuchat, 1987; Nout and Rombouts, 1990). Feng et al. (2007) reported the production of volatiles by R. oligosporus during the fermentation of soybean and barley tempeh. R. oligosporus has also been used to increase the functional properties of foods and reduce the levels of allergenic proteins in buckwheat (Handoyo et al., 2006). As noted above, R. oligosporus appears to be a domesticated fungus. It has rarely, if ever, been reliably reported from sources other than fermented foods. References. Hesseltine (1965); Schipper and Stalpers (1984); Zheng et al. (2007a).

Rhizopus oryzae Went & Prins. Geerl. Fig. 6.16 Rhizopus arrhizus A. Fisch.

On CYA and MEA, colonies filling the whole Petri dish with fine greyish mycelium and small blackish grey sporangia; reverse pale. On G25N, colonies

163

30–60 mm diam, or occasionally filling the whole Petri dish, relatively low and sparse. No growth at 58C. Colonies at 378C covering and sometimes filling the Petri dish, similar to that on CYA or more sparse. Sporangiophores borne in clusters of one to three from rhizoids, with stipes up to 1500 mm long, usually unbranched; sporangia spherical, up to 150 mm diam, white at first then becoming greyish black at maturity; columellae usually spherical, up to 100 mm diam, pale brown, in age often collapsing downwards to form umbrella shapes; sporangiospores brown, of variable shape, ellipsoidal to broadly fusiform or irregularly angular, commonly 5–8 mm long, with striate walls. Distinctive features. This species is distinguished from Rhizopus stolonifer by its smaller sporangia and spores and by much faster growth at 378C. It differs from R. microsporus and R. oligosporus by longer stipes, larger collumellae and larger spores. Taxonomy. Schipper and Stalpers (1984) concluded that Rhizopus oryzae was the correct name for this species, considering R. arrhizus to be a doubtful name, placing this latter species in synonymy with R. oryzae. Their treatment is followed here, as R. oryzae is by far the more common name in the literature. In the latest taxonomy, Zheng et al.

Fig. 6.16 Rhizopus oryzae (a) colonies on CYA, 7 days, 258C; (b) sporangia and columellae, bar ¼ 25 mm; (c) sporangiospores, bar ¼ 10 mm

164

(2007a) have taken up R. arrhizus based on neotypification by Ellis (1985), so that usage of this name may increase in the future. However, it is not clear if Ellis’ neotypification is valid. Physiology. Rhizopus oryzae has been reported to grow from 7 to 428C or 458C, with the optimum near 378C (Panasenko, 1967; Gleason, 1971; Schipper, 1984; Zheng et al., 2007a). Its minimum aw for growth is 0.88 (Hocking and Miscamble, 1995). Growth under optimal conditions is exceptionally fast (up to 1.6 mm/h), although slower than that of R. stolonifer (Hocking and Miscamble, 1995). R. oryzae produces lipase D, an enzyme which hydrolyses triglycerides of fatty acids and acts as a catalyst for the esterification

6 Zygomycetes

of fats and oils to improve their physical properties. The toxicity of lipase D produced by R. oryzae was examined in rats for 13 weeks and no ill effects were detected (Flood and Kondo, 2003). Mycotoxins. Maize meal on which isolates of Rhizopus oryzae had been grown was toxic to ducklings and rats, causing growth depression (Rabie et al., 1985). The toxin responsible has not been elucidated. Ecology. Rhizopus oryzae is an important agent of human zygomycoses, causing mainly rhinocerebral infections (de Hoog et al., 2000). It has been isolated quite frequently from foods, often as R. arrhizus. It appears to be associated with many of the rots in fruits and vegetables usually attributed to R. stolonifer

Fig. 6.17 Rhizopus sexualis (a) colonies on CYA, 7 days, 258C; (b) sporangium, bar ¼ 25 mm; (c) zygospores, bar ¼ 50 mm; (d) sporangiospores, bar ¼ 10 mm

6.6

Genus Syncephalastrum

165

(Snowdon, 1990, 1991). From our observations, it is more common in tropical commodities than is R. stolonifer. It was especially common in peanuts, being isolated from 21% of all peanut kernels examined from Indonesia, 15% of those from Thailand and 14% of those from the Philippines. It was growing in 25% of all copra pieces from Thailand and 9% of kemiri nuts from Indonesia. It was present in lower numbers in maize, various kinds of beans, sorghum and cowpeas (Pitt et al., 1993, 1998a). Other reports include maize (Askun, 2006), peanuts, pecans, hazelnuts, pistachios, wheat, barley, potatoes, sapodillas and various other tropical foods (see Pitt and Hocking, 1997). References. Schipper (1984); de Hoog et al. (2000); Zheng et al. (2007a).

Rhizopus sexualis (G. Sm.) Callen

Fig. 6.17

On CYA, colonies covering the whole Petri dish, very low and sparse, of white to greyish mycelium; sporangia in limited numbers, pale; black zygospores conspicuous; reverse uncoloured. On MEA, colonies similar to that on CYA, though more dense. On G25N, colonies at least 40 mm diam, sometimes covering the whole Petri dish, often as vigorous as on CYA or MEA, and of similar appearance, but zygospores usually absent. No growth at 5 or 378C. Sporangiophores borne from rhizoids, one to three per cluster, stipes up to 1500 mm long, usually unbranched; sporangia spherical, 50–150 mm diam, white, becoming grey; columellae spherical to ellipsoidal, up to 100 mm diam or long, in age collapsing to umbrella shapes; sporangiospores subspheroidal, ellipsoidal or angular, 5–12(–25) mm long, with thick, grey, striate walls. Zygospores commonly occurring, 80–180 mm diam, black, with adjacent cells (suspensors) approximately spherical, but at maturity appearing hemispherical. Distinctive features. Rhizopus sexualis is distinguished from other foodborne species of Rhizopus and Mucor by the profuse production of conspicuous black zygospores. Physiology. Schipper (1984) reported that this species only grows below 308C; however, Zheng et al. (2007a) reported 26–278C as the maximum growth temperature. Relatively fast growth on

G25N perhaps indicates a relatively low minimum aw for growth. Mycotoxins. Mycotoxin production has not been reported. Ecology. Rhizopus sexualis causes a soft rot of strawberries (Harris and Dennis, 1980) and occasionally of other fruit. References. Schipper (1984); Zheng et al. (2007a).

¨ 6.6 Genus Syncephalastrum J. Schrot. Syncephalastrum produces sporangiospores in cylindrical sacs (merosporangia) attached around spherical vesicles, providing a superficial resemblance to Aspergillus niger at low magnifications. In other respects, this genus resembles Mucor. There is one species, S. racemosum.

¨ Syncephalastrum racemosum J. Schrot. Fig. 6.18 On CYA and MEA, colonies covering the whole Petri dish, sparse to grey; reverse pale or yellowish brown. On G25N, colonies 20–30 mm diam, dense to floccose, grey; reverse pale. No growth at 58C. Colonies at 378C filling the whole Petri dish, dark grey; reverse yellow brown. Sporangiophores borne from aerial hyphae, stipes long and branched or produced as short side branches from fertile hyphae; sporangial heads 30–80 mm diam, with sporangiospores formed in a single line within cylindrical sacs (merosporangia) borne on spicules around the vesicle; vesicles spherical or nearly so, 10–50 mm diam, brown, with walls smooth except at merosporangium attachment points, usually collapsing irregularly; sporangiospores adhering in chains of up to 10, becoming brown, irregular in size and shape, spherical to cylindrical, 3.0–5.0(–10) mm diam or long, smooth walled. Distinctive features. See genus description. Physiology. Good growth has been reported between 17 and 408C (Domsch et al., 1980): no doubt growth limits are rather wider than these figures. This is one of the Mucorales most tolerant to low aw: the minimum for growth is 0.84 aw (Hocking and Miscamble, 1995).

166

6 Zygomycetes

Fig. 6.18 Syncephalastrum racemosum (a) colonies on CYA, 7 days, 258C; (b) columellae, bar ¼ 25 mm; (c) stages in merosporangium formation, bar ¼ 25 mm; (d) sporangiospores, bar ¼ 10 mm; (e) developing merosporangium, bar = 10 mm; (f) sporangiospores, bar ¼ 10 mm

Mycotoxins. It was reported by Makun et al. (2007b) that two of three isolates of Syncephalastrum were mildly toxic to mice (one of three injected with extracts died). No other details are known.

Ecology. Reported isolations from foods have not been numerous, but this species is of widespread occurrence nevertheless. Principal reported sources have been from nuts (see Pitt and Hocking, 1997),

6.7

Genus Thamnidium

167

young coconuts (Waje et al., 2005), cereals, spices (Elshafie et al., 2002), fermented foods, processed meats and capsicums (see Pitt and Hocking, 1997). We isolated this species from 10% of all coriander seeds examined from Indonesia and 4% of all kemiri nuts. It was present in 2% of all sorghum grains from Thailand; 1% of Thai mung beans; Philippine maize, soybeans and black pepper; and Indonesian peanuts. It was present at lower levels in soybeans, black and red beans, cassava and copra from Thailand and mung beans, maize and milled rice from Indonesia (Pitt et al., 1994, 1998a). Recent reports include cashews (Adebajo and Diyaolu, 2003), millet (Makun et al., 2007) and chickpeas (Dawar et al., 2007b). References. Benjamin (1959); O’Donnell (1979).

6.7 Genus Thamnidium Link As well as large columellate sporangia, Thamnidium produces sporangioles, small sporangia without columellae, borne on highly branched structures. Sporangioles in Thamnidium contain sporangiospores, similar to those borne in sporangia. There is a single species, T. elegans.

Thamnidium elegans Link

Fig. 6.19

On CYA, colonies usually covering the whole Petri dish, sparse but quite deep due to the production of

long sporangiophores, coloured grey to pale brown; reverse pale. On MEA, colonies 30–50 mm diam, similar to those on CYA but with longer, larger sporangiophores. On G25N, colonies variable, from germination only to colonies 10 mm diam, low and dense. At 58C, colonies 15–35 mm diam, of low mycelium. No growth at 378C. Sporangiophores very large, borne from surface hyphae, stipes bearing sporangia or sporangioles or both; sporangia brown, 150–250 mm diam; columellae roughly spherical, 40–80 mm diam, or sometimes larger, collapsing or tearing irregularly; sporangioles 12–25 mm diam, rough walled; sporangiospores narrowly to broadly ellipsoidal, 6–15 mm long, with thin, smooth walls. Distinctive features. See genus description. Physiology. Thamnidium. elegans is psychrophilic, growing down to at least 18C (Brooks and Hansford, 1923). The maximum temperature for growth is about 278C (Gleason, 1971). Mycotoxins. Mycotoxin production has not been reported. Ecology. Thamnidium elegans has traditionally been associated with cool stored meat, on which it occurs as long ‘‘whiskers’’. Improved temperature control of meat storage has virtually eliminated this problem in developed countries. However, where meat is traditionally hung for curing, festoons of T. elegans can still sometimes be seen. Whether this constitutes ‘‘spoilage’’ depends on definitions. Spoilage of Taleggio

Fig. 6.19 Thamnidium elegans (a) colonies on CYA, 7 days, 258C; (b) sporangiophore, developing sporangioles and sporangiospores, bar ¼ 25 mm

168

cheese (a traditional, washed rind, soft, red pigmented cow’s milk cheese from Italy) has been reported (Dragoni et al., 1997). We isolated T. elegans from coriander from Indonesia, where it

6 Zygomycetes

was found in 1% of seeds examined (Pitt et al., 1998a). References. Hesseltine and Anderson (1956); O’Donnell (1979).

Chapter 7

Penicillium and Related Genera

Grouped in this chapter are genera which produce conidia in a structure termed a penicillus (Latin, little brush). A penicillus consists essentially of a well-defined cluster of phialides or similar cells bearing small, single-celled, dry conidia in chains. The phialides are either attached to a stipe directly or through one or more stages of branching. Branches are of generally similar diameter to stipes (Fig. 7.1). This definition covers four genera here: Penicillium, Geosmithia, Paecilomyces and Scopu-

lariopsis. Genera with greatly enlarged stipe apices, e.g. Aspergillus, are excluded, as are genera in which phialides are not borne in clusters or which produce wet, adherent conidia, e.g. Fusarium, Trichoderma. The degree of relatedness of these four genera treated here is not high, but they are conveniently considered together. A variety of characteristics enable them to be distinguished. They are keyed out here on the basis of differences in phialide shape and conidial shape and colour.

Key to genera producing penicilli 1

Mature conidia truncate or flattened at the base; colonies white, buff or brown Conidia not truncate at the base, symmetrical from end to end; colony colour various

2 (1)

Mature conidia spherical to ellipsoidal, in shades of blue, green and/or grey Mature conidia ellipsoidal to fusiform or cylindroidal, not blue, green or grey

Penicillium 3

3 (2)

Phialides cylindrical, rough walled, necks truncate, mature conidia cylindroidal Phialides gradually tapering, smooth walled, often with long necks angled away from the phialide axis, conidia ellipsoidal to fusiform

Geosmithia

Teleomorphs. The genera in this chapter are all associated with Ascomycete teleomorphs. Treated here are Byssochlamys, Eupenicillium and Talaromyces. Microascus, the teleomorph of Scopulariopsis, does not occur in foods. Byssochlamys is the teleomorph of certain species of Paecilomyces, while Eupenicillium is associated with Penicillium species. Talaromyces teleomorphs are produced by some species of Geosmithia, Paecilomyces

Scopulariopsis 2

Paecilomyces

and Penicillium, but those in Paecilomyces are not found associated with foods. It should be emphasised that production of teleomorphs by any of these three anamorph genera is the exception rather than the rule. Species producing teleomorphs are discussed here under the teleomorph name, as the teleomorph is usually the dominant form or is more significant in foods because of properties such as heat resistance.

Key to teleomorph genera producing anamorphs with penicilli 1

Asci produced within a discrete macroscopic structure, a cleistothecium or gymnothecium Asci produced openly, solitarily or in groups on hyphae

2 Byssochlamys

2 (1)

Asci produced within a cleistothecium, a body with a solid cellular wall Asci produced within a gymnothecium, with walls of fine, closely woven hyphae

Eupenicillium Talaromyces

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_7,  Springer ScienceþBusiness Media, LLC 2009

169

170

7

Penicillium and Related Genera

Fig. 7.1 Scanning electron micrographs of various types of penicilli, showing the relationship between the diameter of the stipe and the other elements of the penicillus

In contrast with the very large genus Penicillium, the other genera considered in this chapter include few species of significance in foods. The smaller genera are considered below in alphabetical order, followed by Penicillium.

7.1 Genus Byssochlamys Westling Byssochlamys has the distinction of being almost uniquely associated with food spoilage, and in particular with the spoilage of heat processed acid foods. Byssochlamys species have been reported only rarely from other sources. Byssochlamys is an Ascomycete genus characterised by the absence of cleistothecia, gymnothecia or other bodies enveloping asci during development. Asci in Byssochlamys are borne in open clusters, in association with, but not surrounded by, unstructured wefts of fine, white hyphae. Techniques for isolating Byssochlamys rely on a heat treatment to inactivate other fungal spores which are less heat resistant. These techniques have been described in Chapter 4.

Three species of Byssochlamys are significant in food spoilage: B. fulva, B. nivea and B. spectabilis. B. nivea is readily distinguished from the others by its white colonies. It also forms ascospores readily in the laboratory, when incubated for 10 days at 25 or 308C (but not 378C). B. fulva and B. spectabilis form brown colonies of quite similar appearance. B. fulva usually forms ascospores in the laboratory when grown at 308C (but not at 25 or 378C). B. spectabilis does not usually make ascospores in laboratory culture, as it is heterothallic, i.e. usually two isolates must be mated to form the teleomorph. For this reason, B. spectabilis is discussed under its anamorph Paecilomyces variotii. How do we know we have a Byssochlamys if it does not produce ascospores? Presumptive evidence of a Byssochlamys can be made from plates at 25 or 378C if the isolate has come from heat processed foods or raw materials. If a brown Paecilomyces is isolated after a sample has been heat treated at 758C, it is probably B. spectabilis, which is included under Paecilomyces variotii later in this chapter. This aspect is discussed further under ‘‘heat resistant fungi’’ in Chapter 12.

Key to Byssochlamys species included here 1

Colonies on CYA and MEA persistently white to cream Colonies on CYA and MEA predominantly buff to brown

2.

White asci and ascospores produced on MEA at 308C after 7–10 days; conidia mostly cylindroidal Colonies remaining buff to brown, no obvious teleomorph; conidia mostly ellipsoidal

B. nivea 2

B. fulva Byssochlamys spectabilis See Paecilomyces variotii

7.1

Genus Byssochlamys

Byssochlamys fulva Olliver & G. Sm.

171

Fig. 7.2

Anamorph: Paecilomyces fulvus Stolk & Samson Colonies on CYA and MEA at least 60 mm diam, often covering the whole Petri dish, relatively sparse, low or somewhat floccose; conidial production heavy, uniformly coloured olive brown or paler; reverse in similar colours or pale. Colonies on G25N 5–10 mm diam, texture variable, low and sparse to deep and floccose, coloured white or as on CYA. No growth at 58C. At 30 and 378C, colonies usually covering the whole Petri dish, low and sparse, coloured as on CYA or brighter; reverse in similar colours. Teleomorph single asci borne from, but not enveloped by, wefts of contorted white hyphae, best developed at 308C on MEA, maturing in 7–12 days, occasionally formed at 258C in fresh isolates but maturing slowly if at all; asci spherical to subspheroidal, 9–12 mm diam; ascospores ellipsoidal, 5–7 mm long, smooth walled. Anamorph reproductive structures penicilli, best observed at 258C, borne from surface hyphae or long, trailing, aerial hyphae; stipes 10–30 mm long; phialides of nonuniform appearance, flask shaped or narrowing gradually to the apices, 12–20 mm long; conidia mostly cylindroidal or doliiform (barrel shaped), usually narrow and 7–10 mm long, but sometimes longer, wider or ellipsoidal from individual phialides, smooth walled. Distinctive features. In culture at 308C, Byssochlamys fulva is distinguished by rapidly growing brown colonies with areas of fine white hyphae, in which asci are produced in open clusters. At 258C, colonies may not produce the white hyphae, and then they closely resemble Paecilomyces variotii (teleomorph Byssochlamys spectabilis). The simplest microscopic distinguishing feature is that P. variotii produces ellipsoidal not cylindroidal conidia. Moreover, P. variotii does not readily produce a teleomorph in culture. Physiology. The major physiological characteristic which makes Byssochlamys fulva significant in food mycology is the heat resistance of its ascospores. First noted by Olliver and Rendle (1934), who described this species, and very carefully documented by Hull (1939), this property has been extensively studied in more recent work. These studies were comprehensively reviewed by Beuchat and Rice (1979).

Many variables can affect the heat resistance of Byssochlamys fulva, which can vary markedly from isolate to isolate (Bayne and Michener, 1979; Hatcher et al., 1979). Factors such as pH, water activity (see Dijksterhuis and Samson, 2006) and the presence of preservatives also have an effect. Ascospores are more susceptible to heat if the pH is low, and/or if preservatives such as sodium benzoate or potassium sorbate are present (El-Geddawy, 2005). On the other hand, high levels of sugar have a protective effect (El-Geddawy, 2005). For B. fulva, a D value of between 1 and 12 min at 90C8 (Bayne and Mitchener, 1979) and a z value of 6–78C (King et al., 1969) are practical working values. Ascospores of B. fulva isolated from canned tomato paste were more heat resistant in tomato juice than in phosphate buffer (Kotzekidou, 1997a). Treatment at 808C for 1 hour had no effect on the viability of ascospores, however, in the presence of CO2, a considerable reduction (80%) in ascospore numbers was observed (Ballestra and Cuq, 1998). Byssochlamys fulva ascospores are also very resistant to high pressure, with resistance increasing with ascospore age (Chapman et al., 2007). After treatment at 600 MPa for 10 min, a 4–5 log reduction was achieved for 3 week old ascospores, whereas the same treatment applied to 9 week old ascospores resulted in a 1-log reduction and 15 week old ascospores were activated after pressure treatment (Chapman et al., 2007). The second physiological characteristic which makes Byssochlamys fulva an outstanding spoilage fungus is its ability to grow at very low oxygen tensions. This ability, shared with B. nivea, but apparently not with other common heat resistant moulds, provides Byssochlamys species with a selective advantage in products such as canned, bottled or cartoned fruits and fruit juices. Our observations suggest that in the presence of very low levels of oxygen, these species grow anaerobically and produce CO2. A small amount of oxygen contained in the headspace of a jar or bottle, or the slow leakage of oxygen through a package such as a Tetra-Brik can provide sufficient oxygen for these fungi to grow. The production of gas may cause visible swelling and spoilage of the product. Byssochlamys fulva can grow extensively in atmospheres containing 20, 40 or 60% CO2 and less than 0.5% O2. In 20% CO2, colonies reached 40–80 mm diameter in 15–30 days, depending on the medium. In 60% CO2 and less than 0.5% O2,

172

7

Penicillium and Related Genera

Fig. 7.2 Byssochlamys fulva (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) asci and ascospores, bar = 5 mm; (e) conidia, bar = 5 mm

7.1

Genus Byssochlamys

colonies of 20–40 mm diameter were formed in 30 days; colonies in 80% CO2 and 20% O2 were similar (Taniwaki, 1995; Taniwaki et al., 2001a). The small amount of growth formed while residual oxygen is consumed in canned or bottled fruit may be sufficient to allow the production of pectinolytic enzymes. The first sign of spoilage by Byssochlamys species is usually a slight softening of the fruit. This progresses until total disintegration takes place, due to the production of powerful pectinases by the fungus (Hull, 1939; Beuchat and Rice, 1979). Off odours, and a slightly sour taste may develop and, as noted above, gas production may occur. It is rare for canned and bottled fruit to be spoiled by fungi other than Byssochlamys species. Mycotoxins. Rice et al. (1977) reported patulin production by some isolates of Byssochlamys fulva, however, there is no published evidence that patulin has been produced by growth of B. fulva in commercially processed fruit juices. Very low levels of patulin were produced under atmospheres containing less than 0.5% O2 and 20–60% CO2 (Taniwaki, 1995). A recent survey revealed that none of the 11 isolates of B. fulva examined produced patulin in vitro (Puel et al., 2007), a result also demonstrated by Houbraken et al. (2006). B. fulva synthesises byssochlamic acid (Houbraken et al., 2006). Ecology. Spoilage caused by Byssochlamys fulva was first recognised in canned strawberries in England in the 1930s. Olliver and Rendle (1934) investigated the incidence of B. fulva in fruits, in other canning ingredients and in packing materials. Although strawberries and plums seemed to be particularly affected, B. fulva was also found on other fruits such as berries, currants and apples (Olliver and Rendle, 1934). Hull (1939) found B. fulva on leaves, fruits and straw from strawberry fields, and also on mummified fruit. Olliver and Rendle (1934) and Hull (1939) concluded that soil acts as the primary reservoir for Byssochlamys ascospores, and that fruit which came in contact with soil directly or from rain splash were susceptible to contamination. Byssochlamys fulva has been reported from several other areas of the world, notably the United States, where it has occurred in grape products (King et al., 1969; Splittstoesser et al., 1971), and Australia, as the cause of spoilage of canned strawberries (Richardson, 1965), fruit juices and fruit-based baby foods. The main Australian source of B. fulva has been shown

173

to be passionfruit juice (Hocking and Pitt, 1984). Effective measures to overcome this problem include washing of fruit before juice extraction, rejection of difficult to clean wrinkled fruit and screening of juices for heat resistant ascospores (Cartwright and Hocking, 1984; Kotzekidou, 1997a). For details of screening methods see Chapter 4. The high heat resistance of Byssochlamys fulva and B. nivea ascospores has led to the study of other approaches for deactivating them. Microwave heating (used to pasteurise packaged ravioli) had little effect on B. nivea (Dragoni et al., 1990). Intermittent use of dimethyldicarbonate was suggested (van der Riet and Pinches, 1991) after it was found that a single high dose (1000 mg/l) had no effect on ascospores of B. fulva (van der Riet et al., 1989). Approximately 1.2 kGy of ionising radiation produced one decimal reduction in viable ascospores of two resistant strains of B. fulva; it was estimated that more than 7 kGy would be needed for an effective pasteurising process. Apple juice containing high numbers of spores and treated with 5 kGy spoiled after 3 months (van der Riet and van der Walt, 1985). References. Samson (1974); Beuchat and Rice (1979); Hocking and Pitt (1984); Houbraken et al. (2006).

Byssochlamys nivea Westling

Fig. 7.3

Anamorph: Paecilomyces niveus Stolk & Samson Colonies on CYA 40–50 mm diam, low and quite sparse, white to slightly grey; reverse pale to mid brown. Colonies on MEA covering the whole Petri dish, low and sparse, white to creamish, with small knots of dense hyphae; reverse pale to brownish. On G25N, usually only microscopic growth. No growth at 58C. At 308C on CYA, colonies covering the whole Petri dish, similar to on MEA at 258C, but often more dense, enveloping distinct knots of dense hyphae. At 378C, colonies 50–70 mm diam, low to floccose, moderately dense, white to cream, reverse pale to brown. Teleomorph similar to that of Byssochlamys fulva except for slightly smaller asci (8–11 mm diam) and ascospores (4–6 mm diam), maturing in 10–14 days at 258C and in 7–10 days at 308C, but rarely found at 378C. Anamorphs of two kinds produced, aleurioconidia and penicilli; aleurioconidia borne singly, common at 30 and 378C, spherical to pyriform, 7–10 mm

174

7

Penicillium and Related Genera

Fig. 7.3 Byssochlamys nivea (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) ascospores, bar = 5 mm; (e) conidia, bar = 5 mm

diam; penicilli sparsely produced, with short stipes bearing irregular metulae and phialides or phialides alone, and with phialides sometimes borne solitarily from hyphae as well; phialides 12–20 mm long, cylindrical then gradually tapering; conidia ellipsoidal to pyriform, 3–6 mm long, smooth walled. Distinctive features. Byssochlamys nivea is readily distinguished from B. fulva by its persistently white

colonies. It differs from other fungi by forming three characteristic types of reproductive structures: aleurioconidia, penicilli and solitary asci. Physiology. The minimum aw for growth of Byssochlamys nivea was 0.92 at 218C, 0.89 at 308C and 0.87 aw at 378C (Roland and Beuchat, 1984). So far as has been established, the physiology of B. nivea is similar to that of B. fulva. Like B. fulva, it

7.2

Genus Eupenicillium

causes spoilage of heat processed acid foods under conditions of low oxygen tension. B. nivea ascospores are reported to be marginally less heat resistant than those of B. fulva (Put and Kruiswijk, 1964; Beuchat and Rice, 1979). A D value at 888C of 0.75–0.8 min, with z values ranging from 4.0–6.1C8, was determined by Casella et al. (1990). See also Engel and Teuber (1991) and Quintavalla and Spotti (1993). B. nivea ascospores exhibited pressure resistance similar to that reported for B. fulva (Butz et al., 1996; Chapman et al., 2007). Remarks about the physiology of Byssochlamys fulva probably apply equally well to B. nivea. Growth of B. nivea in atmospheres containing 0.5% O2 and 20, 40 or 60% CO2 was somewhat slower than observed for B. fulva, while growth on 20% O2 and 80% CO2 was rather faster (Taniwaki, 1995). Taniwaki et al. (2001a) showed that B. nivea was capable of continued growth at 40% CO2 with <0.5% O2. Mycotoxins. Byssochlamys nivea may produce patulin (Rice et al., 1977), byssochlamic acid and mycophenolic acid (Houbraken et al., 2006), but none of these have been reported to be significant in foods. Several studies have investigated the effect of physical parameters on patulin production, including the effect of headspace in glass jars of heat processed grape juice (Rice, 1980), controlled atmospheres (Orth, 1976; Taniwaki, 1995), temperature, aw and preservatives (Roland and Beuchat, 1984; Roland et al., 1984). Minimum aw values for patulin production were 0.92 at 218C and 0.87 at both 30 and 378C (Roland and Beuchat, 1984). Ayesha and Viswanath (2006) reported the production of patulin in sugar cane juice inoculated with B. nivea. Patulin synthesis was greater at 208C than 358C, and more patulin was produced in juice than in commercial broth. Ecology. Byssochlamys nivea appears to be a less common problem in foods than B. fulva. It has mostly been reported from European sources: from sweet cider in Switzerland, from fruit juices in Denmark, from bottled strawberries in the Netherlands and packaged ravioli in Italy (see Pitt and Hocking, 1997). Spoilage of the latter product was prevented by packing in an atmosphere of 60% CO2, 39.4% N2 and 0.05% O2 (Dragoni and Cantoni, 1988). B. nivea has also been isolated from passionfruit juice and from cream cheese in Australia (A.D. Hocking, unpublished). References. Put and Kruiswijk (1964); Samson (1974); Beuchat and Rice (1979); Hocking and Pitt (1984); Houbraken et al. (2006).

175

7.2 Genus Eupenicillium F. Ludw. The name Eupenicillium was applied to the very hard (sclerotioid) cleistothecial state produced by certain Penicillium species as early as 1892. However, for many years it remained common practice to name both teleomorph and anamorph species producing a Penicillium anamorph by their Penicillium name. As well as conflicting with provisions of the International Code of Botanical Nomenclature, this practice ignored the effects of the teleomorph on cultural appearance, longevity, heat and chemical resistance, etc. These important characteristics are readily overlooked if the Penicillium name is used. Benjamin (1955) revived the name Carpenteles Langeron 1922 for Penicillium species producing cleistothecia. This name slowly gained acceptance until Stolk and Scott (1967) showed that the obscure Eupenicillium F. Ludw. 1892 was the correct name for this Ascomycete genus. Scott and Stolk transferred all Carpenteles species to Eupenicillium and described 16 new species (Scott and Stolk, 1967; Stolk and Scott, 1967). Despite vigorous opposition from some taxonomists (Raper, 1957; Fennell, 1973), the use of teleomorph names for Penicillium (and Aspergillus) species where Ascomycete states are present is now firmly established (Pitt, 1979b; Samson and Pitt, 1985; Pitt and Samson, 1993; Pitt and Hocking, 1997; Pitt et al., 2000). Scott (1968) published a monograph of Eupenicillium. Pitt (1974b, 1979b, 2000), while providing new keys and descriptions, adhered closely to Scott’s species concepts. Stolk and Samson (1983) published a revised taxonomy with greatly modified species concepts. Acceptance was limited, as Scott’s concepts appear sound. The close relationships between some Eupenicillium species, some Penicillium species producing sclerotia (cleistothecia which never mature) and indeed some strictly anamorphic species were recognised first by Pitt (1974b). Chemotaxonomy (Frisvad et al., 1990a) and molecular studies (LoBuglio et al., 1993) have added weight to this concept, which probably has phylogenetic significance (Pitt, 1995). Eupenicillium is characterised by the production of macroscopic (100–500 mm diam), smooth walled, often brightly coloured cleistothecia, in association with a Penicillium anamorph. In many species cleistothecia become rock hard as they develop, and may remain so for many weeks or months, finally maturing from the centre to yield numerous eight spored asci. Because of this delayed

176

7

maturation, taxonomy is very difficult if based on the morphology of mature ascospores, as the keys of Scott (1968) and Stolk and Samson (1983) were. For this reason, Pitt (1974b, 1979b) introduced synoptic keys to Eupenicillium, which permit identification of many Eupenicillium species from cultural and microscopic characters. Most Eupenicillium species are soil fungi, and of little interest to the food microbiologist. However, they have occurred from time to time as survivors of heat processing. Williams et al. (1941) recorded that a new species, Penicillium lapidosum (stone-like, an apt name), was causing spoilage of canned blueberries. It possessed highly heat resistant sclerotia (immature cleistothecia). This fungus was later shown to produce a Eupenicillium state. Two points are worth noting: first, it was the immature cleistothecium itself which was acting as the heat resistant body; and second, most Eupenicillium species produce heat resistant ascospores. Fortunately, they find their way into heat processed foods only rarely.

Penicillium and Related Genera

Eupenicillium species have been isolated as heatresistant contaminants of fruit juices on several occasions (Anon., 1967; van der Spuy et al., 1975; Hocking and Pitt, 1984, and unpublished). No particular species appears to be responsible, and growth of the fungus in the product has been rare. As a cause of food spoilage, Eupenicillium ascospores can be safely ignored unless an unusual set of circumstances leads to excessive contamination of some raw material or product. Of the 37 Eupenicillium species recognised by Pitt (1979b), five are treated here: E. brefeldianum, E. cinnamopurpureum, E. hirayamae, E. javanicum and E. ochrosalmoneum. E. cinnamopurpureum and E. javanicum are relatively common. Because Penicillium species that produce sclerotia cannot be distinguished from Eupenicillium species after 7 days incubation, P. raistrickii, P. sclerotiorum, P. simplicissimum and P. thomii are included in this key.

Key to Eupenicillium and sclerotigenic Penicillium species included here 1

Colonies on CYA less than 20 mm diam, coloured deep brown Colonies on CYA more than 20 mm diam, not coloured deep brown

2 (1)

Colonies with bright yellow or orange mycelium and/or reverse colours Colonies with mycelium and reverse uncoloured, or in colours other than bright yellow or orange

E. cinnamopurpureum 2 3 5

3 (2)

Penicilli strictly monoverticillate Penicilli biverticillate or sometimes irregular

4 E. ochrosalmoneum

4 (3)

Growth on CYA at 378C similar to at 258C No growth at 378C

E. hirayamae P. sclerotiorum

5 (2)

Penicilli monoverticillate Penicilli biverticillate

6 (5)

Growth at 378C

6 7 E. javanicum E. brefeldianum P. thomii

No growth at 378C 7 (5)

Usually growth at 378C, conidial walls rough No growth at 378C, conidia with smooth walls

Eupenicillium cinnamopurpureum D.B. Scott & Stolk

Fig. 7.4

Penicillium pusillum G. Sm. Penicillium cinnamopurpureum S. Abe ex Udagawa

Anamorph: Penicillium phoeniceum J.F.H. Beyma Colonies on CYA 15–20 mm diam, of closely textured, sulcate or wrinkled white to brown mycelium, usually

P. simplicissimum P. raistrickii

enveloping numerous brown to pinkish cleistothecia; conidial production sparse, coloured greyish green (25C-D3, 26D2); clear exudate and purple soluble pigment typically produced; reverse pink, purple or cinnamon, rarely pale or dull orange. Colonies on MEA 13–15 mm diam, plane, usually with a central area of cinnamon to brown cleistothecia surrounded by white mycelium and few to numerous penicilli; colours

7.2

Genus Eupenicillium

as on CYA except reverse sometimes dull yellow or orange, usually also with some purple areas. Colonies on G25N 8–12 mm diam, of dense white mycelium, conidial production light to moderate, grey green; reverse purple, pink or cinnamon. At 58C, usually no germination. At 378C, colonies 5–8 mm diam, of white mycelium or, rarely, no growth. Cleistothecia 150–250 mm diam, pinkish cinnamon to brown, becoming hard, maturing very slowly; ascospores ellipsoidal, pale yellow, 3.0–3.5 mm long, with spinulose walls and with two low longitudinal flanges. Conidiophores borne from surface mycelium, stipes 20–150 mm long, strictly monoverticillate, smooth

177

walled, terminating in enlarged (4–5 mm) apices; phialides ampulliform to acerose, 8–12 mm long, gradually tapering; conidia subspheroidal to ellipsoidal, 2.0–3.5 mm long, with smooth or finely roughened walls. Distinctive features. Eupenicillium cinnamopurpureum, together with its anamorph Penicillium phoeniceum, is a distinctive and readily recognised species. It grows slowly on CYA and MEA, stipes are strictly monoverticillate, are apically enlarged and bear long, gradually tapering phialides. Most isolates produce brown cleistothecia, which mature very slowly. Taxonomy. Some isolates fail to produce cleistothecia at all, but are still readily recognisable as

Fig. 7.4 Eupenicillium cinnamopurpureum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) asci and ascospores, bar = 5 mm; (f) conidia, bar = 5 mm

178

7

Penicillium and Related Genera

belonging here. Penicillium phoeniceum is the appropriate name in such cases. Physiology. This species is among the most xerophilic of the Penicillia, growing down to 0.78 aw in glycerol media (Hocking and Pitt, 1979), or 0.82 aw in salt (25% w/v; Udagawa and Tsuruta, 1973). From growth data in Pitt (1979b: 67), this species will grow at temperatures between 4–68C and 35–388C, with some isolate to isolate variation. Mycotoxins. Mycotoxin production has not been reported. Ecology. A very widely distributed species, Eupenicillium cinnamopurpureum has been isolated from flour (Graves and Hesseltine, 1966; as Penicillium sp.), rice (Udagawa, 1959; as P. cinnamopurpureum), dried beans and rice, especially in long term storage (Udagawa and Tsuruta, 1973; Tsuruta and Saito, 1980, as P. pusillum), dried peas (Smith, 1939, as P. pusillum) and stored maize kernels (Wicklow et al., 1998). We isolated this species quite frequently from Southeast Asia: from soybeans, red beans and copra in Thailand, from maize, peanuts, soybeans and black pepper in the Philippines, and from maize, peanuts, kemiri nuts, mung beans, soybeans and red kidney beans in Indonesia. The strict anamorph P. phoeniceum was found in black rice from Thailand and milled rice from Indonesia (Pitt et al., 1994, 1998a). Eupenicillium cinnamopurpureum and Penicillium phoeniceum were amongst the most commonly isolated Penicillia in Australian wheat and various milling fractions (flour, semolina, bran and wheat germ) during a survey of the microbiology of flour milling (Hocking, unpublished). References. Udagawa (1959); Udagawa and Tsuruta (1973); Pitt (1979b, 2000).

(5B6-A8). Colonies on MEA usually 15–22 mm diam, but up to 35 mm if cleistothecia absent, plane, otherwise similar to those on CYA. Colonies on G25N 8–14 mm diam, similar to those on CYA. No germination at 58C. At 378C, colonies 20– 30 mm diam, mycelium centrally brown, otherwise similar to those produced at 258C. Cleistothecia buff to yellow, appearing orange or brown from adherent hyphae, 250–300 mm diam or up to 400 mm long, hard, maturing after 4–6 weeks or more; ascospores small and ellipsoidal, yellow, 2.2–3.0 mm long, with rough walls and two small longitudinal flanges. Conidiophores borne from ropes of aerial hyphae, with stipes 10–50 mm long, smooth walled, strictly monoverticillate; phialides ampulliform, 6–8(–10) mm long; conidia subspheroidal, minute, 1.8–2.8 mm long, smooth walled. Distinctive features. Relatively slowly growing, brilliant yellow and orange coloured colonies make Eupenicillium hirayamae a readily recognised species. Confirmation is provided by the formation of very similar colonies at 25 and 378C and by the production of short monoverticillate conidiophores. Taxonomy. Most isolates produce cleistothecia and in due course ascospores. A few isolates have been encountered which do not – interestingly colony growth is significantly faster. Such isolates are correctly identified as Penicillium hirayamae. Physiology. We are not aware of any physiological studies on this species. Mycotoxins. Mycotoxin production has not been reported. Ecology. Eupenicillium hirayamae has mostly been isolated from cereals, from Thailand, United States South Africa and India. References. Udagawa (1959); Pitt (1979b, 2000).

Eupenicillium hirayamae D.B. Scott & Stolk Fig. 7.5

Eupenicillium javanicum (J.F.H. Beyma) Stolk & D.B. Scott Fig. 7.6

Anamorph: Penicillium hirayamae Udagawa

Penicillium javanicum J.F.H. Beyma (name invalid because it included the teleomorph).

Colonies on CYA usually 22–28 mm diam, but up to 45 mm diam if cleistothecia absent, radially sulcate, of dense, brilliant yellow or orange mycelium usually enmeshing cleistothecia and overlaid by funicles (ropes) of fertile hyphae; conidial production moderate, dull green (28D3); exudate clear to pale yellow; reverse usually apricot to deep orange

Anamorph: Penicillium indonesiae Pitt Colonies on CYA 30–45 mm diam, radially sulcate, consisting of dense, velutinous, pale yellow mycelium; cleistothecia abundant, enveloped by the mycelium; conidia sparse; exudate copious, brown; reverse olive green, often also with deep reddish

7.2

Genus Eupenicillium

179

Fig. 7.5 Eupenicillium hirayamae (a) colonies on CYA and MEA, 7 d, 258C; (b) penicilli, bar = 10 mm; (c) conidia, bar = 5 mm; (d) asci and ascospores, bar = 5 mm

brown areas. Colonies on MEA 30–50 mm diam, similar to those on CYA but mycelium brighter yellow and reverse usually deep olive brown. Colonies on G25N 9–14 mm diam, of floccose, pale yellow mycelium; reverse olive brown. No germination at 58C. At 378C, colonies 25–50 mm diam, radially sulcate or irregularly wrinkled, mycelium usually white but sometimes deep brown; cleistothecia sometimes present but penicilli absent; clear to brown or reddish exudate and reddish brown soluble pigment usually produced; reverse pale, yellow, reddish brown or deep brown. Cleistothecia dull yellow to brown, 80–200 mm diam, maturing in 2–3 weeks; ascospores

ellipsoidal, 2.5–3.0 mm long, with slightly roughened walls and a faint longitudinal furrow. Conidiophores borne from aerial hyphae, stipes 50–100 mm long, smooth walled, nonvesiculate, bearing monoverticillate penicilli; phialides ampulliform, 8–11 mm long; conidia subspheroidal, ellipsoidal or pyriform, 2.5–3.0 mm long. Distinctive features. Eupenicillium javanicum grows rapidly at 378C, produces sparse strictly monoverticillate penicilli and produces bright colours in exudate and reverse on CYA at 258C. Mature ascospores are produced relatively rapidly, usually within 14 days at 258C.

180

7

Penicillium and Related Genera

Fig. 7.6 Eupenicillium javanicum (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) conidia, bar = 5 mm; (e) asci and ascospores, bar = 5 mm

Taxonomy. The anamorph name, necessary for nomenclatorial reasons at the time it was published (Pitt, 1979b), is hardly ever used. Physiology. Tanaka et al. (2000) reported that Eupenicillium javanicum exhibited greater cellulase activity on soy bran medium than on crystalline cellulose medium. Mycotoxins. Some isolates of Eupenicillium javanicum have been reported to produce xanthomegnin and a little palitantin (Frisvad et al., 1990a). This is unlikely to be of practical importance. A report of patulin production (Okeke et al., 1993) requires confirmation, especially as this compound was not found by Frisvad et al. (1990a). Two new compounds, namely eujavanoic acids A and B, were

characterised from E. javanicum. The decalin derivatives were found to be active against Aspergillus fumigatus (Okamoto et al., 2004). Ecology. This is one of the most common Eupenicillium species, abundant in soils, but occurring in foods rather rarely. It has been isolated from wheat and flour (Basu and Mehrotra, 1976), fermented and cured meats (Leistner and Ayres, 1968) and peanuts (Pitt et al., 1998a). Additional species. Eupenicillium brefeldianum (B.O. Dodge) Stolk and D.B. Scott is closely related to E. javanicum; the two species produce colonies of similar sizes on the standard media. E. brefeldianum produces longer, sometimes metulate stipes, larger

7.2

Genus Eupenicillium

181

cleistothecia (150–250 mm long) and ascospores (3–4 mm long) and pale orange mycelium. It lacks green reverse colours. This species has been reported to cause spoilage of apple juice in South Africa by surviving pasteurising treatments (Anon., 1967; van der Spuy et al., 1975). It has also been isolated from peanuts (Joffe, 1969). References. Stolk and Scott (1967); Pitt (1979b, 2000).

Eupenicillium ochrosalmoneum D.B. Scott & Stolk

Fig. 7.7

Anamorph: Penicillium ochrosalmoneum Udagawa

Colonies on CYA 18–28 mm diam, plane or sulcate, with a moderately deep, dense layer of mycelium enveloping abundant cleistothecia, overlaid by sparse to abundant penicilli; mycelium white to yellow (2-4A7); conidia greyish green to dull green (27C-D3); clear to pale yellow exudate and pale to bright yellow soluble pigment sometimes produced; reverse yellow to cadmium orange (3-5A8). Colonies on MEA 15–20 mm diam, plane, low and sparse to moderately deep and dense, velutinous; mycelium white at the margins, elsewhere bright yellow, as on CYA; cleistothecia abundant, on the agar surface; conidial production sparse to moderate, coloured as on CYA; yellow soluble pigment

Fig. 7.7 Eupenicillium ochrosalmoneum (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) conidia, bar = 5 mm; (e) asci and ascospores, bar = 5 mm

182

usually produced; reverse bright yellow to orange. Colonies on G25N 1–7 mm diam, of deep, dense white mycelium; reverse pale or yellow. No germination at 58C. At 378C, colonies 10–30 mm diam, radially sulcate or wrinkled, closely resembling growth on CYA at 258C except for increased cleistothecial production by some isolates. Cleistothecia pale to bright yellow or orange, 250–500 mm diam, sclerotioid, maturing in 4–6 weeks or more; ascospores ellipsoidal, 3.5–5.0 mm long, with walls smooth to roughened or spinulose, ornamented with two longitudinal flanges. Conidiophores borne from surface or aerial hyphae, stipes variable, (20–)40–200 mm long, smooth walled, bearing very irregular penicilli, basically biverticillate but with subterminal and intercalary metulae often present, the latter being indistinguishable from short monoverticillate conidiophores; phialides ampulliform-acerose 6–8(–10) mm long; conidia subspheroidal to apiculate, commonly 2.5– 3.5 mm long, smooth walled, borne in disordered chains. Distinctive features. Eupenicillium ochrosalmoneum forms slowly growing, bright yellow colonies on both CYA and MEA at 258C. Growth on CYA at 378C is similar in size and colouration to that at 258C. These characteristics are shared with E. hirayamae: this latter species, however, produces strictly monoverticillate penicilli. Physiology. Eupenicillium ochrosalmoneum is capable of growth down to 0.88 aw (Hocking and Pitt, 1979). Mycotoxins. This species is the second major producer of citreoviridin, a mycotoxin described in more detail under Penicillium citreonigrum. Citreoviridin produced by E. ochrosalmoneum has been found naturally occurring in US maize, at levels up to 2.8 mg/kg (Wicklow et al., 1988). Ecology. This species has been reported from maize in North America (Wicklow and Cole, 1984; Wicklow et al., 1988) and pig feed in Slovakia (Labuda and Tancˇinova´, 2003). We have seen both Eupenicillium ochrosalmoneum and its anamorph in Australian wheat and wheat fractions during a survey of the microbiological status of Australian flour mills (Hocking, unpublished). Its importance lies in the potential to produce citreoviridin under natural conditions. References. Scott and Stolk (1967); Pitt (1979b, 2000).

7

Penicillium and Related Genera

7.3 Genus Geosmithia Pitt Geosmithia was split from Penicillium by Pitt (1979a), because phialide shape and wall texture, and conidial shape and colour, are distinctive. Conidia in Geosmithia are strictly cylindrical and borne from usually rough walled, cylindrical phialides which have cylindrical necks. Except in one species, G. namyslowskii K.M. Zalessky, known only from a single isolate from soil, conidia totally lack the blue or green pigmentation so characteristic of Penicillium. Recent molecular studies (Ogawa et al., 1997) and later papers have shown that Pitt’s concept is polyphyletic and that some Geosmithia species are closely related to Penicillium. The species treated here, G. putterillii, remains in Geosmithia with the type species, G. lavendula. The main habitat of Geosmithia species, including G. putterillii, is in association with bark beetles (Kolarˇ ı´ k et al., 2004). However, G. putterillii is quite common on cereal products and may cause spoilage occasionally (Fig. 7.8).

Geosmithia putterillii (Thom) Pitt

Fig. 7.8

Penicillium putterillii Thom

Colonies on CYA 25–35 mm diam, velutinous, floccose or somewhat funiculose, mycelium white or buff, conidial production moderate to heavy, off white to buff or very pale yellow; reverse dull yellow or olive brown. Colonies on MEA 20–30 mm diam, usually markedly funiculose, otherwise as on CYA. Colonies on G25N 15–18 mm diam, similar to those on CYA. At 58C, no germination to formation of microcolonies. At 378C, usually no growth, sometimes colonies to 8 mm diam. Conidiophores borne from surface or aerial hyphae, stipes 20–100 mm long, with rough walls; penicilli often complex, with 2, 3 or more branch points; phialides rough walled, cylindrical, 8–10 mm long; conidia cylindrical, 4.0–5.0  2.0–2.5 mm, smooth walled. Distinctive features. Geosmithia putterillii forms quite slowly growing off white to buff colonies, with irregular penicilli and distinctive cylindrical conidia.

7.4

Genus Paecilomyces

183

Fig. 7.8 Geosmithia putterillii (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d, e) penicilli, bars ¼ 10 mm; (f) conidia, bar ¼ 5 mm

Taxonomy. Kolarˇ ı´ k et al. (2004) published a comprehensive molecular and morphological study of Geosmithia putterillii and related taxa. Physiology. Information on the physiology of Geosmithia putterillii is lacking. However, judged by its isolation from a variety of dry substrates, this species is xerophilic. Mycotoxins. Mycotoxin production has not been reported. Ecology. Most foodborne isolations of this species have come from cereals and cereal products, katsuobushi and dried peas (see Pitt and Hocking, 1997), although Kolarˇ ı´ k et al. (2004) report that it is normally associated with bark beetles. In our laboratory, it has twice been recorded as the cause

of spoilage of Lebanese bread, on which it produces a profuse white growth. We have also isolated it from Indonesian soybeans (Pitt et al., 1998a). It has been reported (as Penicillium pallidum) as one cause of rots in melons (Snowdon, 1991). Reference. Pitt (1979a); Kolarˇ ı´ k et al. (2004).

7.4 Genus Paecilomyces Bainier Paecilomyces was split from Penicillium by Bainier (1907a) on the basis of differences in phialide shape and conidial colour. Paecilomyces phialides have necks which are characteristically (although not exclusively) long and bent away from the phialide

184

7

axis. Unlike those produced by Penicillium species, conidia are never green or blue and are usually elongate, fusiform or ellipsoidal to cylindroidal. Paecilomyces species are of importance as soil fungi and insect pathogens. Only three species are commonly isolated from foods: P. variotii and P. lilacinus are treated below. The teleomorph of P. variotii is Byssochlamys spectabilis (Houbraken

Penicillium and Related Genera

et al., 2006). P. variotii is heterothallic, requiring culturing of (+) and (–) strains together to produce the teleomorph, so the Byssochlamys state is rarely seen in culture. The discovery of the teleomorph explains why P. variotii is often seen in heat-treated samples without evidence of a heat-resistant state (Houbraken et al., 2008). P. fulvus is described under its teleomorph, Byssochlamys fulva.

Key to Paecilomyces species included here 1

Colonies on CYA and MEA brown Colonies on CYA and MEA lilac or mauve

2 P. lilacinus

2 (1)

Conidia typically cylindrical; on CYA at 308C, superficial white hyphae at colony centres Conidia ellipsoidal to fusiform; on CYA at 308C, superficial white hyphae absent

Paecilomyces variotii Bainier

Fig. 7.9

Teleomorph: Byssochlamys spectabilis (Udagawa & Suzuki) Samson et al. Talaromyces spectabilis Udagawa & Suzuki

Colonies on CYA of variable size, 30–70 mm diam, plane, of low to floccose appearance, usually coloured uniformly brown or olive brown from conidia; reverse pale. Colonies on MEA 70 mm diam or more, otherwise very similar to those on CYA. Colonies on G25N 8–16 mm diam, similar to on CYA or of white mycelium. No germination at 58C. At 378C, colonies growing very rapidly, 60 mm or more diam, similar to those at 258C, or with sporulation reduced; reverse pale. Penicilli borne from aerial hyphae on short stipes, of irregular pattern, a cluster of phialides alone or with metulae and phialides or occasionally rami; phialides 12–20 mm long, tapering gradually, with collula often bent away from the axis; conidia mostly subspheroidal to ellipsoidal, sometimes cylindroidal or pyriform, usually 3.0–5.0 mm long, smooth walled. Distinctive features. With its rapidly growing colonies, coloured uniformly brown to olive brown at both 25 and 378C, and irregular penicilli with long phialides, Paecilomyces variotii is a readily recognised fungus. Only P. fulvus is similar: P. variotii is distinguishable by its broadly ellipsoidal rather than cylindrical conidia. When grown on MEA at 308C for 10 days, P. fulvus usually produces its Byssochlamys state, whereas isolates of P. variotii

P. fulvus (see Byssochlamys fulva) P. variotii

do not usually produce asci. However, if P. variotii is isolated from a heat treated food or beverage, it is likely to be the anamorph of Byssochlamys spectabilis (Houbraken et al., 2008). Taxonomy. Houbraken et al. (2006) reported that Byssochlamys spectabilis is the teleomorph of Paecilomyces variotii. The teleomorphic state is rarely seen in laboratory culture because B. spectabilis is heterothallic: production of ascospores requires co-culturing of (+) and (–) strains (Houbraken, et al., 2008). For a description of the teleomorphic state, see Udagawa and Suzuki (1994). Physiology. Paecilomyces variotii is a xerophilic fungus, with growth recorded down to 0.80 aw (Wheeler and Hocking, 1988). It grows between about 5 and 45–488C, with an optimum at 35–408C (Samson, 1974). Our unpublished experiments indicate an ability to grow under low oxygen tensions. Mycotoxins. Houbraken et al. (2006) reported viriditoxin production by Byssochlamys spectabilis. Ecology. This species is a ubiquitous contaminant of foods and raw materials. It has a strong association with edible oils, raw materials containing oil, bacon, peanuts and peanut cake (see Pitt and Hocking, 1997), margarine (Demirci and Arici, 2006) and cocoa beans (Wolcik-Stopczynska, 2006). Amongst many other reports, it has been recorded from cereals (Loiveke et al., 2004), bread, meat products, biltong, health foods and airline foods (see Pitt and Hocking, 1997). In our laboratory, we have frequently isolated Paecilomyces variotii from heat processed fruit

7.4

Genus Paecilomyces

185

Fig. 7.9 Paecilomyces variotii (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

products and beverages, including sports drinks. It was found in 7% of all black peppercorns we examined from Indonesia (Pitt et al., 1998a), and at low levels in black beans, cassava, cashews and copra from Thailand (Pitt et al., 1994). Paecilomyces variotii is notable for sorbate resistance and causes spoilage of margarine and processed cheeses due to production of 1,3-pentadiene from the degradation of sorbate (Sensidoni et al., 1994). It is also one cause of spoilage in dried fruits and other products. This is due to the ‘‘mouldy leather’’ odour of chloroanisole produced from chlorophenols (Tindale et al., 1989; Whitfield et al., 1991).

References. Samson (1974); Udagawa and Suzuki (1994); Houbraken et al. (2006, 2008).

Paecilomyces lilacinus (Thom) Samson

Fig. 7.10

Penicillium lilacinum Thom

Colonies on CYA 25–35 mm diam, plane, dense to floccose; mycelium in marginal areas white, elsewhere pale pink or pinkish grey, sometimes also areas of bright pale yellow; conidial production sparse, pinkish brown; reverse pale, yellow or brown. Colonies on MEA 25–32 mm diam, low

186

and sparse, mycelium uniformly pinkish grey; reverse pale or centrally brown. Colonies on G25N 3–6 mm diam, low and dense. No germination at 58C. At 378C, response ranging from no growth to colonies up to 5 mm diam. Conidiophores borne from aerial or surface hyphae, 200–600 mm long, with finely roughened

7

Penicillium and Related Genera

walls, bearing irregular verticils of metulae both terminally and subterminally; phialides 7–10  2.5–3.0 mm, tapering to long, narrow collula; conidia ellipsoidal to fusiform, 2.5–3.0  2.0–2.2 mm, with smooth to finely roughened walls. Distinctive features. This is the only pink species of Paecilomyces at all common in foods.

Fig. 7.10 Paecilomyces lilacinus (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d, e) penicilli, bars = 10 mm; (f) conidia, bar = 5 mm

7.5

Genus Scopulariopsis

Physiology. Paecilomyces lilacinus grows from 8 to 388C and between pH 2 and 10 in submerged culture (Duncan, 1973). It is capable of growth down to at least 0.90 aw in NaCl (Tresner and Hayes, 1971) or carbohydrate media (Panasenko, 1967). Mycotoxins. Paecilotoxin is the only reported mycotoxin (Mikami et al., 1989). Its significance is unknown. Khan et al. (2003) recently tested one strain of Paecilomyces lilacinus for the production of paecilotoxin and were unable to show toxin production in that strain, suggesting that toxin synthesis may vary among isolates (Khan et al., 2003; Park et al., 2004b). Ecology. Although a contaminant rather than a food spoilage fungus, Paecilomyces lilacinus has been isolated from a variety of foods: cereals, including wheat, flour and barley, peanuts and pecans, fresh figs, beans, frozen meat, salami (see Pitt and Hocking, 1997) and in our laboratory from bottled water and pandanus nuts from Papua New Guinea. Reference. Samson (1974).

7.5 Genus Scopulariopsis Bainier Like Paecilomyces, Scopulariopsis was segregated from Penicillium by Bainier (1907b); it is generally agreed that the differences from Penicillium are more fundamental. In Scopulariopsis, conidia are not extruded from phialides, but are cut off from annelides. The difference from a phialide is clearly evident: as each successive conidium is cut off from the tip of an annelide, a small amount of wall material remains, so that the annelide elongates, and in age shows a succession of faint rings or scars. A consequence is that conidia borne from annelides possess distinctive broad bases. In Scopulariopsis, this characteristic is readily seen under the microscope at high magnification. Colonies of Scopulariopsis species range in colour from white to brown, but are never green or blue. Most species are broad ranging saprophytes, of common occurrence in decaying vegetation and soil. In comparison with their ubiquity in these habitats, they must be classed as relatively uncommon in foods. Only one species, S. brevicaulis, warrants description here.

187

Scopulariopsis brevicaulis (Sacc.) Bainier Fig. 7.11 Colonies on CYA usually 40–50 mm diam, low, dense and velutinous, often irregularly wrinkled, coloured orange grey to brownish orange (5B2-C4); reverse bright yellow to orange brown. Colonies on MEA usually 40–50 mm diam, sometimes much smaller, 15–30 mm, low and sparse at the margins, sometimes centrally floccose, plane, coloured brownish orange (6C4-5); reverse yellow brown. Colonies on G25N 15–20 mm diam, low and dense, white or centrally yellow, reverse white to bright yellow. No growth at 58C. At 378C, colonies 7–20 mm diam, dense, often centrally raised, white or brown; reverse dull yellow. Reproductive structures varying from single conidiogenous cells (annelides) to irregular penicilli, sometimes with well defined metulae and rami; conidia pyriform with a distinctly truncate base, clearly visible before release from the annelide, 5–8 mm diam, brown, with rough walls, sometimes adhering in short chains. Distinctive features. Small single-celled conidia with truncate bases are characteristic of Scopulariopsis. Colonies of S. brevicaulis at 258C are low, dense and brown. Growth at 378C is slow. The brown, rough walled conidia are distinctive. Physiology. The minimum aw for growth of this species is 0.90 (Galloway, 1935). Better growth was observed in media supplemented with 0.1 to 1 M NaCl (Yoder et al., 2003). Mycotoxins. Mycotoxins are not produced. Ecology. Inagaki (1962) isolated Scopulariopsis brevicaulis from rice grains and rice flour, nonfat dried milk and butter. This species was reported to be a cause of spoilage in cheese (Northolt et al., 1980), along with other rarer species of Scopulariopsis. Other sources have been barley, wheat, salami, matured cheese, bacon and biltong (see Pitt and Hocking, 1997) and luncheon meats (Mohamed and Hussein, 2004). We have isolated this species from soybeans and black pepper from the Philippines, and peanuts and mung beans from Indonesia (Pitt et al., 1998a). Reference. Morton and Smith (1963).

188

7

Penicillium and Related Genera

Fig. 7.11 Scopulariopsis brevicaulis (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) conidia, bar = 5 mm

7.6 Genus Talaromyces C.R. Benj. The name Talaromyces is derived from the Greek word for a basket, which aptly describes the body in which asci are produced by this Ascomycete genus. Known as a gymnothecium, this ascocarp is composed of fine hyphae woven into a more or less closed structure of indeterminate size.

Talaromyces is characterised by the production of yellow or white gymnothecia in association with an anamorph characteristic of Penicillium, Paecilomyces or Geosmithia. The production of gymnothecia in association with other anamorphs characterises a variety of other genera, unknown in foods and outside the scope of this book.

7.6

Genus Talaromyces

189

As with Eupenicillium, until recently Talaromyces species with Penicillium anamorphs were commonly known by their Penicillium names. It is current practice for use of the anamorph names to be restricted to references to the anamorphs alone. Talaromyces is a genus of about 25 species, mostly soil inhabiting. Their main interest to the food mycologist lies in their production of heat resistant ascospores and, consequently, their isolation from pasteurised fruit juices and fruit-based

products (Hocking and Pitt, 1984). The most commonly isolated heat-resistant species is T. macrosporus, but we have also seen T. trachyspermus a number of times. The most common Talaromyces species in nature is T. flavus, which is occasionally isolated from commodities such as cereals. T. wortmannii, readily distinguished from T. flavus by its slower growth but similar in many other respects, is the only other species likely to be encountered in foods.

Key to Talaromyces species included here 1

Colonies exceeding 25 mm diam on MEA in 7 days Colonies not exceeding 25 mm diam on MEA in 7 days

2 (1)

Ascospores more than 5 mm long Ascospores less than 5 mm long

3 (1)

Colonies yellow Colonies white

¨ Talaromyces flavus (Klocker) Stolk & Samson

2 3 T. macrosporus T. flavus T. wortmannii (see T. flavus) T. trachyspermus

Fig. 7.12

Penicillium vermiculatum P.A. Dang. (invalid name, teleomorph was included) Talaromyces vermiculatus (P.A. Dang.) C.R. Benj.

Anamorph: Penicillium dangeardii Pitt

Colonies on CYA 18–30 mm diam, occasionally larger, plane, low and quite sparse to moderately deep and floccose; mycelium bright yellow, less commonly buff or reddish brown, in most isolates concealing developing gymnothecia; conidial production usually sparse and inconspicuous, but if more profuse, greenish grey; clear to reddish exudate present occasionally; reverse sometimes yellow, more usually orange, reddish or brown. Colonies on MEA 30–50 mm diam, generally similar to those on CYA but gymnothecia more abundant; reverse usually dull orange or brown, but sometimes deep brown or deep red. Colonies on G25N 2–7 mm diam, low, of sparse white mycelium; occasionally microcolonies or no growth. No germination at 58C. At 378C, colonies 20–45 mm diam, usually similar to those on CYA, but sometimes with white or brown mycelium or overlaid with grey conidia; gymnothecia absent; reverse yellow, orange or brown.

Gymnothecia of tightly interwoven mycelium, bright yellow, about 200–500 mm diam, closely packed, maturing within 2 weeks; ascospores yellow, ellipsoidal, 4.0–5.0 mm long, with spinose walls. Conidiophores borne from aerial hyphae, stipes 20– 80 mm long, bearing terminal biverticillate or less commonly monoverticillate penicilli; phialides acerose, 10–16 mm long; conidia ellipsoidal to fusiform, 2.5–4.0 mm long, with smooth to spinulose walls. Distinctive features. Talaromyces flavus and the closely related T. macrosporus grow relatively rapidly on MEA, produce bright yellow colonies at both 25 and 378C, and usually produce abundant yellow gymnothecia, especially on MEA. T. flavus produces ascospores less than 5 mm long, smaller than those of T. macrosporus. Physiology. Beuchat (1988) and Quintavalla and Spotti (1993) reported that the heat resistance of Talaromyces flavus ascospores (in both cases using strain FRR 1265) was substantially lower than that of isolates with larger ascospores now known to belong to T. macrosporus. No other studies have been published on the physiology of this species. Mycotoxins. This species is not known to produce mycotoxins. Ecology. This is by far the most common species of Talaromyces found universally in soils in warmer climates. Consequently it occurs in foods from time to time, most probably as a contaminant rather

190

7

Penicillium and Related Genera

Fig. 7.12 Talaromyces flavus (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) conidia, bar = 5 mm; (e) ascus and ascospores, bar = 5 mm

than as a spoilage fungus. Reports of T. flavus as a heat-resistant fungus have always referred to T. macrosporus so far as can be established. Additional species. Talaromyces wortmannii (Klocker) Stolk and Samson (anamorph Penicillium ¨ kloeckeri Pitt; synonym P. wortmannii Klocker) ¨ resembles T. flavus closely in colony colours, appearance, gymnothecia, ascospores and penicilli. However, the two species are readily distinguished by differences in growth rates: T. wortmannii produces colonies only 10–15 mm and 15–20 mm diam on

CYA and MEA, respectively, at 258C; at 378C, colonies are less than 10 mm diam. Hocking and Pitt (1979) reported the minimum aw for growth of T. wortmannii as 0.88 at 258C. Isolates of Talaromyces wortmannii produce the mycotoxin rugulosin (Frisvad et al., 1990b) (see Penicillium rugulosum), but this is unlikely to be of practical importance. Like Talaromyces flavus, T. wortmannii is a soil fungus. It is widespread, but relatively uncommon. It sometimes occurs in foods, e.g. wheat, pecans and salami (see

7.6

Genus Talaromyces

191

Pitt and Hocking, 1997). There are no reports of spoilage, or of occurrence in pasteurised products. References. Stolk and Samson (1972); Pitt (1979b).

Talaromyces macrosporus (Stolk & Samson) Frisvad et al.

Fig. 7.13

Talaromyces flavus var. macrosporus Stolk & Samson

Anamorph: Penicillium macrosporum Frisvad et al. Colonies on CYA of variable size, usually 20–40 mm diam, plane, floccose; mycelium pale to bright yellow (3A2-4); immature gymnothecia sometimes present, bright yellow (2-3A6); conidial production sparse and inconspicuous; exudate clear or more commonly red; soluble pigment usually present, orange red (8A6-8); reverse brown to dark brown, or, in the presence of soluble pigment, dark red approaching black. Colonies on MEA usually 45–55 mm diam, sometimes smaller, plane, usually floccose, mycelium pale yellow (3A2-4), usually enveloping abundant developing gymnothecia, bright yellow (2-3A6); exudate and soluble pigment absent; reverse often strongly coloured, orange, brown, red or olive. On G25N, microcolonies to colonies 7 mm diam formed. No germination at 58C. At 378C, colonies usually 40–50 mm diam, occasionally less, plane, floccose, mycelium white to pale yellow; gymnothecia not produced; red exudate and soluble pigment sometimes present; reverse pale, brown, or in the presence of soluble pigment, dark red. Gymnothecia borne as a contiguous layer on the agar surface, or sometimes discrete, 1.0–1.5 mm diam, composed of fine, bright yellow hyphae, maturing within 2 weeks; ascospores ellipsoidal, commonly 5.0–6.5 mm long, with spinose walls. Conidiophores borne from aerial hyphae, sparsely produced, stipes 20–80 mm long, bearing terminal biverticillate or less commonly monoverticillate penicilli; phialides acerose, 10–16 mm long; conidia ellipsoidal to fusiform, 2.5–4.0 mm long, with smooth to spinulose walls. Distinctive features. Like the closely related Talaromyces flavus, T. macrosporus grows relatively rapidly on MEA, produces bright yellow colonies at both 25 and 378C, and usually forms abundant yellow gymnothecia. T. macrosporus produces ascospores that are more than 5 mm long.

Taxonomy. Established for strains of Talaromyces flavus with large ascospores (Stolk and Samson, 1972), T. flavus var. macrosporus was not recognised by Pitt (1979b). On the basis of differences in secondary metabolite production, Frisvad et al. (1990b) raised the variety to species status. Moreover, the quite frequent recovery of isolates with large ascospores from heat-processed foods (Hocking and Pitt, 1984) indicates a real physiological difference from isolates with small ascospores. T. macrosporus is recognised here for isolates with ascospores more than 5 mm long. Physiology. From the food industry viewpoint, the main attribute of Talaromyces macrosporus is the very high heat resistance of its ascospores. Published decimal reduction times and z values vary: a D88 of 7–22 min, a D91 of 2.9–5.4 min (Beuchat, 1986), a D90 of 2–7 min and a z value of 10.3C8 (King and Halbrook, 1987), a D90 of 2.2 min and a z value of 5.2C8 (Scott and Bernard, 1987), a D80 of 190 min and a D90 of 6 min and a z value of 6.7C8 (King and Whitehand, 1990). Dijksterhuis and Teunissen (2004) demonstrated a D85 of 30–100 min. These values are considerably higher than reported for Byssochlamys species (Beuchat, 1986). The influence of organic acids, including preservatives, on the heat resistance of T. flavus was reported by Beuchat (1988). Ascospores of Talaromyces macrosporus contain trehalose, which is degraded to glucose following heat activation. Upon germination, glucose is released from the ascospores. The presence of trehalose in ascospores is thought to act as a stress protectant against dehydration and heat (Dijksterhuis et al., 2002). The effect of sugars such as trehalose in protecting vital enzymes from heat has been shown in other organisms such as Neurospora crassa (Dijksterhuis et al., 2002). Dijksterhuis and Teunissen (2004) showed that high pressure treatments at 400–800 MPa activated ascospores to germinate. The authors explained that high pressure constituted a stress which induced germination, in a situation akin to heat (Dijksterhuis et al., 2002). Ascospore age also affects heat resistance, with mature ascospores being more resistant to heat. This phenomenon has also been shown in ascospore suspensions, where young ascospores stored in buffer developed increased heat resistance over time (Dijksterhuis and Teunissen, 2004). Mycotoxins. This species has been reported to produce duclauxin, a secondary metabolite of

192

7

Penicillium and Related Genera

Fig. 7.13 Talaromyces macrosporus (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) ascospores, bar = 5 mm; (f) conidia, bar = 5 mm

doubtful significance as a mycotoxin (Frisvad et al., 1990a). Ecology. Talaromyces macrosporus (sometimes reported as T. flavus) has been reported fairly frequently from heat processed juices (see Pitt and Hocking, 1997), often as a cause of spoilage. T. macrosporus has rarely been reported away from heat processed juices. It has been found in low levels in pasteurised dairy products (Aydin et al., 2005). References. Stolk and Samson (1972); Frisvad et al. (1990a).

Talaromyces trachyspermus (Shear) Stolk & Samson

Fig. 7.14

Talaromyces spiculosporus (Lehman) C. R. Benj.

Anamorph: Penicillium lehmannii Pitt Colonies on CYA 15–20 mm diam, occasionally smaller, plane, sparse at the margins, otherwise floccose; mycelium white; gymnothecia not produced; conidial production sparse; reverse pale to dull yellow brown. Colonies on MEA 20–25 mm diam, sparse at the margins but floccose and more

7.6

Genus Talaromyces

dense centrally; mycelium white to creamish; gymnothecia tardily produced, few if any conidial structures, reverse pale to deep orange brown. On G25N, no growth or microcolonies. No germination at 58C. At 378C, colonies 25–40 mm diam, sparse at the margins, but deeper and floccose centrally; mycelium white, usually enveloping developing gymnothecia; conidial formation sparse or absent; reverse pale to dull brown. Gymnothecia of densely woven hyphae, cream to pale yellow when mature (2–3 weeks); 300–500 mm diam; ascospores uncoloured, ellipsoidal, 3.5–4.5 mm

193

long, with spinose walls. Conidiophores borne from aerial hyphae, short stipes (5–20 mm long) bearing terminal penicilli, biverticillate or monoverticillate, sometimes with subterminal elements; phialides acerose, 8–15 mm long, conidia variable, ellipsoidal to fusiform, 2.5–4.5 mm long, with smooth walls. Distinctive features. Talaromyces trachyspermus colonies are slow growing, pale, with straw coloured gymnothecia produced readily at 30–378C after 7–14 days. Physiology. Ascospores of Talaromyces trachyspermus are heat resistant. T. trachyspermus was isolated

Fig. 7.14 Talaromyces trachyspermus (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (e) ascospores, bar = 5 mm; (f) conidia, bar = 5 mm

194

from pineapple concentrate heated at 808C for 30 min (Enigl et al., 1993) and we have isolated it from a teabased beverage that had received a heat treatment of 948C for 30 sec. Jesenska´ et al. (1992) isolated T. trachyspermus from soil that had been heated at 708C for 60 min, but not from samples that were heated at 808C for 60 min. The D80 value for T. trachyspermus is probably in the order of 5–10 min with a z value of 2.6 – 0.2C8 (A.D. King, pers. comm.). Mycotoxins. No mycotoxins have been reported from Talaromyces trachyspermus. Ecology. The natural habitat for Talaromyces trachyspermus is soil. It has been reported from soil in Nepal (Minoura et al., 1975), India (Rajak et al., 1991) and the Slovak Republic (Jesenska´ et al., 1992). Isolates in the CBS catalogue have come from soil in Morocco, Nigeria and Germany and bird dung in Venezuela. Food sources include pineapple juice and concentrate (Enigl et al., 1993), canned strawberries, pasteurised drinking yoghurt and honey tea with ginsing (CBS catalogue). In our laboratory all isolates of T. trachyspermus have come from heat processed products: fruit juice jelly, pineapple slices, canned strawberries and an iced tea beverage. References. Stolk and Samson (1972); Pitt (1979a).

7.7 Genus Penicillium Link While it is arguable whether Aspergillus or Penicillium is of greater economic importance as a cause of food spoilage, it is certain that Penicillium is the more diverse genus, in terms of numbers of species and range of habitats. Penicillium species are ubiquitous, opportunistic saprophytes. Nutritionally, they are supremely undemanding, being able to grow in almost any environment with a sprinkling of mineral salts, any but the most complex forms of organic carbon and a wide range of physico-chemical environments, i.e. aw, temperature, pH and redox potential. A majority of the described species are soil fungi, and their occurrence in foods is more or less accidental and rarely of consequence. However, quite a number of species are closely associated with human food supplies. Some species are more specialised: several are destructive pathogens on fruit (e.g. Penicillium digitatum, P. expansum, P. italicum); a few grow

7

Penicillium and Related Genera

below 0.80 aw (e.g. P. brevicompactum, P. chrysogenum, P. implicatum), at low oxygen tension (e.g. P. roqueforti) or are preservative resistant (P. roqueforti). Many are psychrotrophic and capable of causing food spoilage at refrigeration temperatures. A problem with attempting to establish the incidence of Penicillium species in foods is that so many accounts have not identified Penicillia to species level; as a result many surveys of Penicillia in foods are of little value in the present context. Reports on the occurrence of Penicillium species in this book have therefore relied on a relative handful of papers which have provided detailed species lists. Given the taxonomic difficulties of the genus, it is unlikely that these reports are entirely accurate, but in our view they provide the best information available. Taxonomy. Penicillium taxonomy has evolved rapidly since the last complete taxonomy (Pitt, 1979b) was published. Details of these developments may be found elsewhere (Pitt, 1989a; Pitt and Samson, 1990a, b, 1993); the important result is that consensus has been reached on the names and circumscriptions of most species. Publication of a list of ‘‘Names in Current Use (NCU)’’ means that names in Penicillium, Aspergillus and related genera now enjoy protection from earlier names under the International Code of Botanical Nomenclature (Pitt and Samson, 1993; Greuter et al., 1994: p. x). In consequence, the taxonomy of Penicillium has now become quite stable. Fortunately, all of these more recent changes have had little effect on the names of well known species. None of the Penicillium names used either in the previous edition of this book (Pitt and Hocking, 1997) or in the current taxonomy of common species (Pitt, 2000) have required change. However, the taxonomy of the subgenus Penicillium has recently been revised by Samson and Frisvad (2004). The number of species in this subgenus has been increased from about 25 to over 50. Penicillium taxonomy is not easy for the inexperienced. The species commonly occurring in foods are mostly similar in colour and general colony appearance. Reproductive structures are small and often ephemeral. However, it is the authors’ belief that identification of a high percentage of isolates to species level can be accomplished if isolates are grown under standardised conditions of medium and temperature and examined after a relatively short time (seven days), so that fruiting structures

7.7

Genus Penicillium

and colony colours are at their best. Colony diameters are readily measured and provide very valuable information. The standard conditions used throughout this book were originally developed specifically for Penicillium taxonomy. The cultural conditions used and the general principles of colony examination, etc., have been outlined in Chapter 4. As with the other genera described in this chapter, the reproductive structure characteristic of Penicillium is a conidiophore with a relatively delicate, distinct stipe terminating in a penicillus. The form of the penicillus determines the primary taxonomic division of the genus into subgenera. To determine the subgenus to which a Penicillium isolate belongs, count the number of branch points between phialide (or conidial chain) and stipe, down the main axis of the penicillus. In the simplest subgenus, Aspergilloides, there is one branch point: conidia are borne from phialides (the primary conidiogenous cell), which in turn are borne directly from the stipe as a single whorl or verticil (Fig. 7.15a, b). Such penicilli are termed monoverticillate.

195

In its most complex form, the penicillus has characteristically three branch points (terverticillate), and in some species not infrequently four branch points (quaterverticillate), between phialide and stipe (Fig. 7.15c). Species with such penicilli are classified in subgenus Penicillium. As a useful check, nearly all the commonly encountered species in this subgenus characteristically grow to 18 mm in diameter or more on G25N medium in 7 days at 258C. Of intermediate complexity are biverticillate penicilli, i.e. ones having two branch points. In this category, there are two distinct subgenera, Furcatum and Biverticillium (Fig. 7.15d, e, f). Until familiarity breeds confidence, distinguish these two subgenera by observing the following characters: the relative lengths of phialides and their supporting cells (metulae); the number of metulae per stipe; colony diameters on G25N; and if all else fails, the shape of the phialides. The differences between these two subgenera are shown in Table 7.1.

Fig. 7.15 Penicillus types in Penicillium species (a, b) monoverticillate; (c) terverticillate; (d, e) biverticillate, subgenus Furcatum; (f) biverticillate, subgenus Biverticillium; bar = 10 mm

196

7

Penicillium and Related Genera

Table 7.1 Characters distinguishing subgenus Furcatum from subgenus Biverticillium Ratio of metula length to phialide length Metulae per stipe Colony diam on G25N Phialide shape

Subgenus Furcatum

Subgenus Biverticillium

Much greater than one

Approximately one (1–1.2)

Not exceeding five 9–18 mm Flask shaped, gradually tapering to neck (ampulliform)

Usually exceeding five Less than 10 mm Parallel sided, abruptly tapering to neck(acerose)

Because of the size and complexity of the genus, no general key to Penicillium is given. Taxonomy is

simplified by providing a key to subgenera here, and then keys to species in the preamble to each subgenus.

Key to subgenera of Penicillium 1

Penicilli monoverticillate or with only a minor proportion bearing metulae Penicilli commonly biverticillate or more complex

2 (1)

Penicilli predominantly biverticillate or irregularly monoverticillate and biverticillate; colonies on G25N rarely exceeding 18 mm diam Penicilli predominantly terverticillate; colonies on G25N not usually less than 18 mm diam

3 (2)

Subgenus Aspergilloides 2

Penicilli biverticillate or, much less often, terverticillate; ratio of metula to phialide length near one; colonies on G25N less than 10 mm diam Penicilli biverticillate or irregularly monoverticillate and biverticillate; ratio of metula to phialide length much greater than one; colonies on G25N more than 9 mm diam (see also Table 7.1)

7.7.1 Penicillium subgenus Aspergilloides Dierckx In this subgenus conidiophores are strictly or predominantly monoverticillate, i.e. with phialides borne directly on the stipe, with only one branch point between stipe and conidial chain. In some species, metulae may be present on a proportion of stipes (less than 50%). The distinction between species considered to produce monoverticillate or biverticillate penicilli is not always obvious. Some species produce penicilli on hyphal branches which can be interpreted either as short monoverticillate stipes or as metulae. The distinction made here is that if such a hypha, as well as producing intercalary branches, terminally produces a cluster of two or more branches at an

3 Subgenus Penicillium Subgenus Biverticillium Subgenus Furcatum

acute angle, these are interpreted as metulae, the penicillus as biverticillate, and the species is placed in subgenus Furcatum. Where the hypha gives rise only to branches at right angles along its length and neither hypha nor branches have terminal clusters of metulae, the hypha is interpreted as a fertile hypha, each branch as a stipe, the penicilli as monoverticillate, and the species is classified in subgenus Aspergilloides. One character used to differentiate species in subgenus Aspergilloides, but not elsewhere in the genus, is the presence or absence of terminal swellings (vesicles) on the stipes. A stipe is considered to be vesiculate when the terminal swelling is twice the stipe diameter or more. If in doubt, examine colonies grown on MEA, as vesicles are sometimes more obvious on this medium.

Key to Penicillium subgenus Aspergilloides species included here 1

2 (1)

Conspicuous orange, yellow, brown or purple mycelium, soluble pigment or reverse colours on CYA Colonies on CYA lacking bright colours Colonies on CYA coloured brown and/or purple Colonies on CYA coloured orange or yellow

2 5 P. phoeniceum (see Eupen. cinnamopurpureum) 3

7.7

3 (2) 4 (3)

Genus Penicillium

197

Stipes usually vesiculate Stipes not or rarely vesiculate

P. sclerotiorum

4

Colonies at 378C more than 15 mm diam

P. hirayamae (see Eupen. hirayamae)

Colonies at 378C less than 15 mm diam or growth absent 5 (1)

Colonies on CYA greater than 30 mm diam Colonies on CYA not greater than 30 mm diam

6 (5)

Stipes rough walled Stipes smooth walled

P. citreonigrum

6 8 P. thomii

7

7 (6)

Growth at 378C No growth at 378C

P. decumbens P. glabrum P. spinulosum

8 (5)

Stipes usually vesiculate Stipes usually nonvesiculate

P. implicatum P. restrictum

Penicillium citreonigrum Dierckx

Fig. 7.16

Penicillium citreoviride Biourge Penicillium toxicarium L. Miyake (invalid name)

Colonies on CYA 20–28 mm diam, radially sulcate and often centrally wrinkled, dense and velutinous; mycelium white to bright yellow; conidia sparse to moderately abundant, greenish grey (27C-D2); exudate present only rarely, clear to pink; yellow soluble pigment typically produced; reverse usually brilliant yellow, occasionally yellow brown. Colonies on MEA 22–26 mm diam, plane to lightly sulcate, low, dense and velutinous; mycelium white, becoming yellow or buff centrally; conidial production moderate, in colours similar to those on CYA or slightly bluish; exudate produced rarely, clear to yellow; brown soluble pigment sometimes produced; reverse pale, brown or deep reddish brown. Colonies on G25N 11–14 mm diam, coloured similarly to those on CYA; reverse pale to brilliant yellow or brown. At 58C, germination to formation of microcolonies. At 378C, typically no growth, occasionally colonies up to 10 mm diam formed. Conidiophores borne from floccose or less commonly funiculose aerial hyphae, stipes slim and delicate, 60–100 mm long, smooth walled, nonvesiculate, monoverticillate, occasionally with two metulae; phialides ampulliform, length varying with isolate, 5–12 mm long; conidia spherical or near, l.8–2.8 mm diam, with walls smooth to very finely roughened, borne in short disordered chains. Distinctive features. Penicillium citreonigrum produces compact yellow colonies which at most grow

weakly at 5 and 378C; stipes are slender, not apically enlarged; conidia are tiny and smooth walled. Taxonomy. Raper and Thom (1949) used the name Penicillium citreoviride for this species. Pitt (1979b) took up the earlier valid name P. citreonigrum, and this has been accepted. Physiology. The physiology of Penicillium citreonigrum has been little studied. Cardinal temperatures are below 58C, 20–248C and 37–388C (ICMSF, 1996b). Ji et al. (2007) reported an optimum growth temperature of 27–308C for P. citreonigrum (as P. citreoviride). They recorded growth at 0.80 aw at 20, 25 and 308C, but no growth occurred at 158C at aw values from 0.80 to 0.90. However, the authors allowed only 24 h for equilibration of the rice cakes (1.5 cm thick) from an initial aw of 0.92, and as the actual aw did not appear to have been measured, we would consider their data on effect of aw on growth to be unreliable. The minimum aw for growth is not known, but this species is probably a xerophile. Mycotoxins. Penicillium citreonigrum is the major source of citreoviridin (El-Banna et al., 1987b), the cause of the Oriental disease known in Japan as acute cardiac beriberi. Recognised for the past three centuries (Ueno and Ueno, 1972), this disease frequently occurred in young healthy adults, and death could occur within a few days. The work of Sakaki in the 1890s (Ueno and Ueno, 1972) implicated mouldy ‘‘yellow rice’’ as a probable cause and led to a ban on the sale of yellow rice in Japan in 1910. The disease in Japan is now only of historical interest. Uraguchi (1969) and Ueno and Ueno (1972) showed that acute cardiac

198

7

Penicillium and Related Genera

Fig. 7.16 Penicillium citreonigrum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d, e) penicilli, bars = 10 mm; (f) conidia, bar = 5 mm

beriberi was a mycotoxicosis due to the growth of P. citreonigrum in rice. Citreoviridin is produced from 10 to 378C, with a maximum near 208C (Ueno, 1972). Also produced by Eupenicillium ochrosalmoneum, citreoviridin is an unusual molecule consisting of a lactone ring conjugated to a furan ring, with a molecular weight of 402 (Cole and Cox, 1981). It is a neurotoxin, acutely toxic to mice, with intraperitoneal and oral LD50s of 7.5 and 20 mg/kg, respectively (Ueno and Ueno, 1972). Ecology. Although not a commonly isolated species, Penicillium citreonigrum is very widely

distributed. Its occurrence as a cause of spoilage of rice in Japan from time to time has been well documented. P. citreonigrum has been reported from other cereals by several authors, including Graves and Hesseltine (1966), Saito et al. (1971b) and Basu and Mehrotra (1976). It was isolated very occasionally in our laboratory during a survey of the microbiology of Australia wheat and flour milling (Hocking, unpublished). It has been isolated much less frequently from other foods: mung beans (Pitt et al., 1994), spices (Takatori et al., 1977) and jam (Udagawa et al., 1977). Magnoli et al. (2005)

7.7

Genus Penicillium

199

isolated this species from pig feed, which consisted of 60% corn and also from rabbit feed in which corn was replaced by oats, alfalfa and barley. References. Uraguchi (1971); Pitt (1979b, 2000).

Penicillium decumbens Thom

Fig. 7.17

Colonies on CYA commonly 20–30 mm diam, occasionally 40 mm, low and dense, velutinous to lightly floccose; mycelium white to cream; conidial production light to moderate, greyish green to dull

green (25C-D3); reverse pale, dull yellow brown or olive. Colonies on MEA 25–40 mm diam, usually plane, typically low and relatively sparse, less commonly floccose; moderate numbers of conidia produced, coloured as on CYA. Colonies on G25N 11–16 mm diam, usually rather sparse, in general terms similar to those on CYA. At 58C, germination by a proportion of conidia up to formation of microcolonies. At 378C, colonies 5–20 mm diam, velutinous to floccose, coloured white to grey green; reverse pale or brownish.

Fig. 7.17 Penicillium decumbens (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

200

Conidiophores borne from aerial hyphae, stipes short, 20–60(–100) mm long, with thin, smooth walls, monoverticillate, nonvesiculate to somewhat enlarged; phialides ampulliform, long and slender, 8–11(–14) mm long; conidia ellipsoidal, in some isolates also pyriform, smooth walled, 2.5–3.0(–4.0) mm long, borne in short, loose columns. Distinctive features. Penicillium decumbens produces short, somewhat apically swollen, monoverticillate conidiophores from aerial hyphae, and smooth walled, distinctly ellipsoidal, dull green conidia. Colonies usually do not show any other colours. This is one of the few Penicillium species which typically grows at both 5 and 378C. Physiology. In the only recorded physiological study, Penicillium decumbens grew down to 0.87 but not 0.82 aw at 20 and 308C (Valero et al., 2007a). Mycotoxins. No mycotoxins are known. Ecology. A ubiquitous fungus, Penicillium decumbens has been isolated from a wide variety of foods. These include dried peas and beans (see Pitt and Hocking, 1997); several kinds of nuts (Pitt et al., 1998a); flour milling equipment (Hocking, unpublished), flour, rice, infant cereal preparations, soybeans, meat products and fresh vegetables (see Pitt and Hocking, 1997). This fungus was also isolated from Spanish grapes and dried vine fruits, although spoilage was not recorded (Valero et al., 2007a). Spoilage of food by this species appears to be unusual. References. Pitt (1979b, 2000).

Penicillium glabrum (Wehmer) Westling Fig. 7.18 Penicillium frequentans Westling

Colonies on CYA 40–50 mm diam, low, plane to radially sulcate; surface texture strictly velutinous; white mycelium sometimes visible at the centres; conidial production moderate to heavy, dull green to dark green (26-27E-F3-5); exudate produced centrally by some isolates, clear, yellow or brown; bright yellow soluble pigment sometimes produced; reverse varying from pale or greenish to bright yellow, deep orange, brown or occasionally reddish. Colonies on MEA 40–55 mm diam, low, strictly velutinous, plane or centrally umbonate; white mycelium visible at the centres only; conidial production moderate, coloured as on CYA;

7

Penicillium and Related Genera

yellow soluble pigment sometimes produced; reverse sometimes pale, but more commonly strongly coloured, greenish, olive, yellow or brown. Colonies on G25N 17–24 mm diam, sulcate or wrinkled, velutinous; mycelium white. At 58C, at least germination; usually microcolonies or colonies up to 4 mm diam formed. No growth at 378C (Fig. 7.18). Conidiophores borne from subsurface or surface mycelium or to a limited extent from aerial hyphae, stipes (25–)50–100(–200) mm long, with walls smooth to finely roughened, monoverticillate or occasionally with two metulae, commonly vesiculate up to 6 mm diam; phialides numerous, ampulliform, 8–12 mm long; conidia spherical to subspheroidal, 3.0–3.5 mm diam, with walls smooth or very finely roughened, borne in long well-defined columns. Distinctive features. Penicillium glabrum grows rapidly at 258C, producing dark green colonies with a strictly velutinous texture. Stipes are vesiculate; conidia are smooth walled and borne in long columns. This species is distinguishable from P. spinulosum only with difficulty: P. glabrum produces conidia with smooth to finely roughened walls, while those of P. spinulosum are finely roughened to distinctly spinulose (Pitt et al., 1990). Taxonomy. The relationship of Penicillium glabrum with P. spinulosum and some less common species was considered in detail by Pitt et al. (1990). It was agreed that these two species were distinct, but very closely related. Physiology. Penicillium glabrum is a psychrotroph, able to grow down to at least 08C (Mislivec and Tuite, 1970b). Maximum temperatures for growth are near 308C, not, as reported by Domsch et al. (1980), above 408C. As it is closely related to P. spinulosum, P. glabrum is probably a xerophile. Mycotoxins. Mycotoxins are not produced. Ecology. Penicillium glabrum has caused spoilage of cheese and margarine (see Pitt and Hocking, 1997). It has been isolated frequently from a wide range of foodstuffs, in earlier literature under the name P. frequentans: from dried and concentrated products such as maize, peanuts, rice and jam; from fermented and cured meats; from fresh cabbage and yams (see Pitt and Hocking, 1997); from wheat, flour and other milling fractions (Hocking, unpublished) and canned carbonated beverage (our data and Ancasi et al., 2006). We have isolated P. glabrum

7.7

Genus Penicillium

201

Fig. 7.18 Penicillium glabrum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

from bottled water, sports beverages, fruit juices, fruit pure´es and a variety of spoiled dairy products. P. glabrum has also been reported from chestnuts (Overy et al., 2003) and grapes (Serra et al., 2006). Additional species. Penicillium spinulosum Thom closely resembles P. glabrum, the main difference being that P. spinulosum produces spinose conidia (Pitt et al., 1990). P. spinulosum is a xerophile, germinating down to 0.80 aw at 22–258C (Pelhate, 1968; Hocking and Pitt, 1979). Like P. glabrum, this is a psychrotroph and has not been reported to produce mycotoxins. P. spinulosum has been

reported most frequently from wheat and flour and meat products (see Pitt and Hocking, 1997). We have isolated it from soybeans and peanuts in Southeast Asia (Pitt et al., 1998a). References. Pitt (1979b, 2000); Domsch et al. (1980).

Penicillium implicatum Biourge

Fig. 7.19

Colonies on CYA and MEA growing slowly, 15–20 mm diam or less, radially sulcate, strictly velutinous; mycelium low and dense, white or buff;

202

conidial production light to heavy, greyish green to dull green (25-26D-E4-5); pale to deep brown exudate sometimes produced on CYA; brown soluble pigment typically produced; reverse yellow, brown or reddish. Colonies on G25N 8–12 mm diam, velutinous; colours similar to those on CYA except reverse pale, olive or brown. At 58C, usually no germination. At 378C colonies of 5–10 mm diam usually produced, dense and velutinous; reverse brown to deep brown.

7

Penicillium and Related Genera

Conidiophores borne from subsurface or surface hyphae, stipes 30–100 mm long, with walls thin and smooth, monoverticillate, sometimes with two metulae, usually vesiculate up to 4–5 mm diam, but not exclusively so; phialides 8–11 mm long, slim, with short collula; conidia ellipsoidal to subspheroidal, 2.5–3.0 mm long, with thick, smooth or finely roughened walls, borne in loose columns. Distinctive features. Penicillium implicatum produces slowly growing, dense colonies on the

Fig. 7.19 Penicillium implicatum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

7.7

Genus Penicillium

standard media; typical isolates grow slowly at 378C. Conidiophores are relatively short and usually vesiculate, conidia are ellipsoidal and smooth or nearly so. Physiology. This species is one of the most xerophilic of the Penicillia, being able to germinate at 0.78 aw in 10 days (Hocking and Pitt, 1979). Conidiogenesis was unaffected by water activity (0.90 and 0.82) when grown on sterile maize grains, but fungal biomass increased at higher aw (Cahagnier et al., 1993). Mycotoxins. Mycotoxin production has not been reported. Ecology. Perhaps because of its slow growth and xerophilic nature, Penicillium implicatum is readily overlooked in surveys. Pitt (1979b) regarded its basic habitat as soil and Nesci et al. (2006) have reported this species in agricultural soils where maize was cultivated. While not of common occurrence, it is a significant biodeteriogen (Raper and Thom, 1949: 203) and spoilage fungus in dried foods. Doupnik and Bell (1971) reported P. implicatum from spoiled pecans; it was also common in Indonesian kemiri nuts (1% of all nuts examined; Pitt et al., 1998a) and was found at lower levels in Thai cashews and Indonesian peanuts (Pitt et al., 1993, 1998a). It has been isolated mainly from cereals: maize, rice, wheat and flour (see Pitt and Hocking, 1997). It was relatively common in Australian flour and other milling fractions (Hocking, unpublished). It has also been reported from cashews and dried peas, meat products and frozen fruit pastries (see Pitt and Hocking, 1997). References. Pitt (1979b, 2000).

Penicillium restrictum J.C. Gilman & E.V. Abbott Fig. 7.20 Colonies on CYA 18–25 mm diam, plane or lightly radially sulcate, typically deep and floccose; mycelium white; conidia absent or sparsely produced, pale grey to bluish grey; clear exudate occasionally present; reverse pale or light brown. Colonies on MEA 15–25 mm diam, plane or umbonate, texture variable, deeply floccose with sparse conidial production to low and funiculose with abundant conidia; mycelium white; conidia greenish grey; reverse pale to brown. On G25N, colonies 11–14 mm diam,

203

plane, deeply floccose or less commonly mucoid; mycelium white; reverse pale. At 58C, typically no germination; rarely a proportion of conidia with germ tubes. At 378C, typically colonies 5–10 mm diam produced, sulcate or wrinkled, white or grey; rarely no growth. Conidiophores borne from loose aerial hyphae or on MEA sometimes from rudimentary funicles, stipes very short, mostly 10–30 mm long, narrow, smooth walled, strictly monoverticillate, nonvesiculate; phialides ampulliform, short, (4–)6–7 mm; conidia spheroidal, ellipsoidal or less commonly pyriform, 2.0–3.0 mm long, finely to coarsely roughened, borne in short irregular chains. Distinctive features. Penicillium restrictum grows relatively slowly on CYA and MEA at 258C, and slowly at 378C. Colonies are usually floccose, with white mycelium and sparsely produced grey conidia. Penicilli are very small. Physiology. A minimum aw for growth of 0.82 has been reported (Hocking and Pitt, 1979). Mycotoxins. This species does not produce mycotoxins. Ecology. Penicillium restrictum is a soil fungus and has not been reported to cause food spoilage. Wheat and flour are the foods from which it has been most frequently isolated (Kurata and Ichinoe, 1967; Basu and Mehrotra, 1976). Other reports have been from processed meats (Lofti et al., 1983), soybeans (Pitt et al., 1994) and poultry feeds (Magnoli et al., 1998). References. Pitt (1979b, 2000).

Penicillium sclerotiorum J.F.H. Beyma Fig. 7.21 Colonies on CYA and MEA typically 30–40 mm diam, less commonly only 20–25 mm, wrinkled, low to moderately deep, dense, consisting of a layer of mycelium, white at the margins, becoming yellow to brilliant orange nearer the centres, usually enveloping abundant sclerotia and overlaid by scattered penicilli; conidia sparsely produced, greyish turquoise or greenish grey (24-26D2-3); exudate limited to abundant, pale, yellow, orange or orange red; yellow or brown soluble pigment usually produced; reverse orange yellow or orange to coffee coloured on CYA, on MEA similar or orange red. Colonies on

204

7

Penicillium and Related Genera

Fig. 7.20 Penicillium restrictum (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) conidia, bar = 5 mm

G25N 13–18 mm diam, dense and wrinkled, generally in colours similar to those described above. Usually no germination at 58C; occasionally germination observed. No growth at 378C. Sclerotia usually present in fresh isolates, pale, spherical or irregular, 200–400 mm long. Conidiophores borne from surface or subsurface hyphae, stipes 100–300 mm long, slender, with thin, smooth to finely roughened walls, strictly monoverticillate, vesiculate, 4–6 mm diam; phialides numerous, ampulliform, 7–9(–11) mm long; conidia ellipsoidal,

2.5–3.0 mm long, with smooth to finely roughened walls, borne in long and well-defined or irregular columns. Distinctive features. Vivid orange to red colony colours, in both obverse and reverse, are striking characteristics which distinguish Penicillium sclerotiorum from other monoverticillate Penicillium species. Physiology. Judged from its occurrence on a wide range of dried foods, Penicillium sclerotiorum is probably a xerophile. No physiological studies have been reported, however.

7.7

Genus Penicillium

205

Fig. 7.21 Penicillium sclerotiorum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

Mycotoxins. Mycotoxin production has not been reported. Ecology. With the exception of frozen fruit pastries (Kuehn and Gunderson, 1963), this species has been isolated from dried foods: wheat and flour; rice, maize, soybeans and jam (see Pitt and Hocking, 1997). It is not a common species, but we isolated it from peanuts, mung beans and soybeans in Southeast Asia (Pitt et al., 1998a). In the earlier literature, Penicillium sclerotiorum is often known as P. multicolor, a name not accepted by Pitt (1979b). References. Pitt (1979b, 2000).

Penicillium thomii Maire

Fig. 7.22

Colonies on CYA 40–60 mm diam, radially sulcate, often lightly floccose, with white mycelium usually surrounding pale to pinkish brown sclerotia, in central areas overlaid by penicilli borne on long stipes; conidial production moderate, dull green (27D-E3); exudate abundant, clear; reverse buff to yellow, or centrally brown in sclerotigenic isolates. Colonies on MEA 40–55 mm diam, plane or centrally wrinkled, usually floccose; sclerotia sometimes present, borne in a layer near the colony

206

centre, coloured apricot (5B6) or paler; other characteristics similar to those on CYA, except reverse orange brown. Colonies on G25N usually 20–24 mm diam, radially sulcate or wrinkled, low and dense; mycelium white; conidia dark green; reverse buff, olive or brown. Germination always occurring at 58C, usually colonies of 2–4 mm diam formed. No growth at 378C. Sclerotia produced by most isolates, ellipsoidal to irregular in shape, usually 250–350 mm long, rapidly becoming hard, pale at first, then pinkish brown, becoming apricot (5B6) on MEA. Conidiophores borne from surface or aerial hyphae, stipes 200–400 mm long, rough walled, strictly monoverticillate, vesiculate on CYA, less so on MEA;

7

Penicillium and Related Genera

phialides large and crowded, 9–12 mm long, with long narrow collula; conidia ellipsoidal, commonly 3.5–4.0 mm long, finely to coarsely roughened, borne in long, rather irregular columns. Distinctive features. Most isolates of Penicillium thomii produce large sclerotia, coloured a distinctive apricot or salmon shade. Colonies grow very rapidly at 258C; stipes are long, vesiculate and rough walled; conidia are ellipsoidal and also rough walled. Physiology. Judged from quite common occurrence in dried foods, rapid growth on G25N, and affinity with Penicillium glabrum, P. thomii is probably a xerophile. It grows rapidly at 58C. Some isolates of P. thomii recovered from rotten grapes

Fig. 7.22 Penicillium thomii (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

7.7

Genus Penicillium

207

in French vineyards were able to synthesise 2-octen1ol and 2-methylisoborneol, compounds responsible for mushroom and earthy smells, respectively, on malt and grape juice agar (La Guerche et al., 2006). Mycotoxins. Mycotoxin production has not been reported for this species. Ecology. Pitt (1979b) reported that Penicillium thomii is a common fungus, widespread in decaying materials as well as soil. In foods, most isolations have been reported from cereals, especially wheat and barley (see Pitt and Hocking, 1997). It occurred in almost 10% of Indonesian dried fish samples examined by Wheeler et al. (1986) and has also been isolated from meat products, peanuts, pistachios, miso (see Pitt and Hocking, 1997) and grapes (Bau et al., 2005). Under the names P. parallelosporum and P. yezoense, Sasaki reported this species as the cause of spoilage of butter (Pitt, 1979b: 186). We isolated P. thomii from spoiled Australian faba beans and at low levels from peanuts in Thailand and Indonesia (Pitt et al., 1993; 1998a). References. Pitt (1979b, 2000).

7.7.2 Penicillium subgenus Furcatum Pitt Penicillium subgenus Furcatum includes species which produce regularly or irregularly biverticillate penicilli, usually with 2–5 terminal metulae. Species in subgenus Biverticillium also produce biverticillate penicilli, but the two subgenera are distinguishable by several

features. Species in subgenus Furcatum have phialides that are ampulliform, or at least have wide apical pores, and are distinctly shorter than their supporting metulae. Colonies on G25N always exceed 9 mm diam in 7 days at 258C. A few species in this subgenus produce metulae in verticils of 5–9 (like species in subgenus Biverticillium), but are readily recognised as members of subgenus Furcatum by their relatively rapid growth on G25N and by a ratio of metula to phialide length greater than one. Two quite different types of penicillus occur in subgenus Furcatum. Some species produce penicilli in which metulae are almost exclusively borne terminally, i.e. the penicilli characteristically consist of verticils of metulae (Fig. 7.15e). In contrast, a second group of species produces irregularly biverticillate penicilli, with metulae borne terminally, subterminally and lower down on the stipe (Fig. 7.15d). Most isolates classifiable in subgenus Furcatum can be readily assigned to one or other of these two groups, the recognition of which will greatly aid identification. In the key which follows, species have first been separated on penicillus morphology and then on growth rates. Most species in subgenus Furcatum are soil fungi and are of relatively uncommon occurrence in foods. Some are included here because they are sometimes isolated from foods enumerated by dilution plating. However, their occurrence in foods is usually only as ubiquitous contaminants. There are some important exceptions of the species described here: Penicillium citrinum, P. corylophilum, P. fellutanum and P. oxalicum are important in spoilage of one kind of food or another.

Key to Penicillium subgenus Furcatum species included here 1

Penicilli predominantly terminal verticils of metulae Penicilli mostly irregular

2 7

2 (1)

Colonies on CYA at 258C exceeding 35 mm diam Colonies on CYA at 258C not exceeding 35 mm diam

3 5

3 (2)

Stipe walls usually rough Stipe walls smooth

4 (3)

No growth at 378C, conidia 3 mm or less in diameter, smooth walled Usually growth at 378C, conidia up to 4 mm in diameter, rough walled

5 (2)

Penicilli with verticils of 5–8 metulae Penicilli with no more than 5 metulae

4 P. oxalicum P. raistrickii P. novae-zeelandiae P. simplicissimum P. paxilli 6

208

6 (5)

7 (1)

8 (7)

7

Colonies on MEA more than 25 mm diam; metulae often of unequal length Colonies on MEA less than 25 mm diam; metulae of equal length Growth at 378C No growth at 378C

P. corylophilum P. citrinum P. janthinellum

8

Stipes and conidia smooth walled Stipes or conidia rough walled

Penicillium citrinum Thom

Penicillium and Related Genera

Fig. 7.23

Colonies on CYA 25–30 mm diam, radially sulcate, marginal areas velutinous, sometimes floccose centrally; mycelium white in peripheral areas, at the centres white to greyish orange (6B5-6); conidial production moderate, greyish turquoise (24C2-3); exudate clear, pale yellow or pale brown to reddish brown, only rarely absent; soluble pigment bright yellow or absent; reverse yellow, yellow brown, reddish brown or olive. Colonies on MEA 14–18 mm diam, rarely 22 mm, plane or radially sulcate, mycelium white to greyish orange (6B5-6); conidial formation moderate to heavy, grey blue at the margins, elsewhere dull green (26-27E3); reverse pale brown to deep yellow brown. Colonies on G25N 13– 18 mm diam, radially sulcate, velutinous or sometimes floccose centrally; mycelium white; conidia often abundant, dull green; reverse pale, dull brown, yellow brown or olive. No germination at 58C. At 378C, colonies usually 8–10 mm diam of wrinkled white mycelium only. Conidiophores borne from subsurface or surface hyphae, stipes 100–300 mm long, smooth walled, characteristically terminating in well defined verticils of 3–5 divergent metulae, less commonly with a divergent ramus, or metulae produced subterminally or along the stipe; metulae usually of uniform length, commonly apically enlarged or vesiculate, phialides ampulliform, 7–8(–12) mm long; conidia spherical to subspheroidal, 2.2–3.0 mm diam, with walls smooth or very finely roughened, typically borne in long well defined columns, one per metula, arranged in a characteristic whorl on each conidiophore. Distinctive features. Penicillium citrinum is readily recognised by its penicilli, which consist of 3–5 divergent and usually vesiculate metulae, bearing long well defined columns of conidia. Colonies on CYA at 258C are sometimes dominated by copious clear to yellow or brown exudate at the centres. On MEA growth is slower and usually dense, with heavy conidial production.

P. fellutanum P. waksmanii P. janczewskii P. canescens

Physiology. This is a mesophilic species, with the minimum temperature for growth 58C or slightly above, the maximum just above 378C (Pitt, 1973) and the optimum 26–308C (Domsch et al., 1980). At 258C, the minimum aw for growth has been reported as 0.80–0.84 (Galloway, 1935; Pitt and Christian, 1968; Hocking and Pitt, 1979). Penicillium citrinum grows over the pH range 2–10 (Wheeler et al., 1991). Mycotoxins. Penicillium citrinum is the main producer of citrinin, a mycotoxin of moderate toxicity. Citrinin is a significant renal toxin to monogastric domestic animals, including pigs (Keblys et al., 2004) and dogs (Kogika et al., 1993; Krejci et al., 1996). The oral LD50 in mice is about 110 mg/kg (Scott, 1977). Domestic birds are also susceptible: citrinin causes watery diarrhoea, increased food consumption and reduced weight gain due to kidney degeneration in chickens (Mehdi et al., 1981; Glahn et al., 1989), ducklings and turkeys (Mehdi et al., 1984). Citrinin has also been reported to cause teratogenic effects in rats (Singh et al., 2007). An impact of citrinin on human T cells was recently demonstrated (Wichmann et al., 2002, Tammer et al., 2007); however, the overall effect of citrinin on humans remains undocumented. Penicillium citrinum produced citrinin in dry cured ham after 4 days incubation at 208C, compared with only 2 days on YES medium (Bailly et al., 2005). After 16 days, significantly higher amounts of citrinin were produced on ham (1330 mg/kg) than on YES medium (86.9 mg/kg). However, pure citrinin extract added to the surface of cured ham was quickly lost or degraded, with more than 50% lost after only 6 h of incubation at 208C and less than 15% remaining after 192 h (Bailly et al., 2005). Ecology. Penicillium citrinum has been isolated from nearly every kind of food which has been surveyed for fungi. The most common sources are cereals, e.g. rice (Park et al., 2005a; Aziz et al.,

7.7

Genus Penicillium

209

Fig. 7.23 Penicillium citrinum (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) conidia, bar = 5 mm

2006), wheat and barley (Aziz et al., 2006), maize (Mazzani et al., 2004; Aziz et al., 2006), and milled grains and flour (Ogundare and Adetuyi, 2003; Hocking, unpublished; and also see Pitt and Hocking, 1997). Among other reported sources are nuts, blackgram in India, amaranth seeds in Argentina, cinnamon and soy sauce (see Pitt and Hocking, 1997). Penicillium citrinum has also been reported in fermented and cured meats (Tabuc et al., 2004), hams (Bailly et al., 2005), cocoa pulp (Ardhana and Fleet, 2003), young coconuts (Waje et al., 2005), soybeans (Aziz et al., 2006), wine grapes (Bau et al., 2005) and dried vine fruits (Romero et al., 2005), curd and cheese (Kumaresan et al., 2003),

bottled mineral water (Cabral and Ferna´ndez Pinto, 2002) as well as coffee beans, dried beans and peppercorns (see Pitt and Hocking, 1997). In Southeast Asia, Penicillium citrinum was isolated from maize, peanuts, copra soybeans, sorghum and cashews (Pitt et al., 1993, 1994). P. citrinum was very widespread in Indonesian food commodities: we isolated it from dried fish, sorghum, peanuts and kemiri nuts, pepper, coriander, maize and mung beans. Citrinin levels were never significant, however (see Pitt and Hocking, 1997). Instances of food spoilage caused by Penicillium citrinum are rare, but this species is much more than a mere contaminant. Because of its mesophilic

210

7

nature, distribution is world wide and, in addition, its ability to grow down to 0.80 aw helps to secure this species a niche in a very wide range of habitats. References. Pitt (1979b, 2000); Domsch et al. (1980).

Penicillium corylophilum Dierckx

Fig. 7.24

Colonies on CYA 25–35 mm diam, plane to deeply radially sulcate, low, moderately dense, strictly velutinous; mycelium white or rarely buff; conidial production light to moderate, dull green

Penicillium and Related Genera

(25C-E3-4); clear exudate sometimes present; reverse pale, brownish or sometimes centrally dark grey. Colonies on MEA 30–45 mm diam, plane, low and relatively sparse, strictly velutinous; mycelium white or buff; conidia in moderate numbers, dull green (26-27D-E3); clear exudate occasionally present; reverse pale at the margins, but usually dull green to very dark green centrally. Colonies on G25N 10–16 mm diam, plane or centrally wrinkled, dense; mycelium white; colours similar to those on CYA; reverse pale. At 58C, usually germination of conidia

Fig. 7.24 Penicillium corylophilum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d, e) penicilli, bars = 10 mm; (f) conidia, bar = 5 mm

7.7

Genus Penicillium

to formation of microcolonies; occasionally small macroscopic colonies produced. No growth at 378C. Conidiophores borne from subsurface hyphae, stipes 100–250 mm long, smooth walled; on CYA, penicilli usually verticils of 2–5 metulae, but sometimes with subterminal metulae or occasionally a ramus; on MEA usually less complex and frequently monoverticillate; in penicilli with two metulae the offset one often longer than the axial; phialides ampulliform, 7–11 mm long; conidia spherical to subspheroidal, commonly 2.5–3.0 mm diam, smooth walled, borne on CYA in disordered chains, on MEA sometimes in long columns. Distinctive features. Relatively long divergent metulae often of unequal length are characteristic of Penicillium corylophilum. This species produces strictly velutinous colonies; on MEA growth is rapid and usually dark green in reverse. Physiology. This is a xerophilic fungus: Hocking and Pitt (1979) reported germination at 0.80 aw after 38 days at 258C. Growth was observed at 0.85 aw (Guynot et al., 2005). The addition of sorbic acid to fermented bakery products completely inhibited the growth of this fungus regardless of water activity (Guynot et al., 2005). Mycotoxins. Mycotoxins are not known to be produced. Ecology. Penicillium corylophilum has been reported to cause spoilage of high-fat foods, including rapeseed and rapeseed oil (Magan et al., 1993) and margarine (Hocking, 1994). In our experience, P. corylophilum occasionally causes spoilage of low aw foods such as jams. It has been reported quite frequently from cereals: barley, paddy rice, wheat and flour (see Pitt and Hocking, 1997). It has also been isolated from salami (Cantoni et al., 2007), laban rayeb, a fermented dairy product (Ahmed and Abdel-Sater, 2003), peanuts, pecans, hazelnuts, soybeans and frozen fruit pastries (see Pitt and Hocking, 1997). In our laboratory, we have isolated P. coryophilum from a range of substrates including bottled water, fruit juices, thickened cream, cosmetics and petroleum products. P. corylophilum occurred at low levels in mung beans and soybeans in Thailand (Pitt et al., 1994), maize in Indonesia (Pitt et al., 1998a), and peanuts in the Philippines (our unpublished data) References. Pitt (1979b, 2000); Domsch et al. (1980).

211

Penicillium fellutanum Biourge

Fig. 7.25

Colonies on CYA 17–24 mm diam, very dense, radially sulcate, velutinous; mycelium white, usually visible only at the margins; formation of conidia light to heavy, coloured pale grey if sparse, but more commonly dark green (27-28F4); colourless exudate sometimes present; reverse pale. Colonies on MEA 14–18 mm diam, low and dense, radially sulcate, usually velutinous with a floccose central area, less commonly entirely floccose; conidia moderately abundant; wrinkled; colours similar to colonies on CYA. Colonies on G25N usually 12–16 mm diam, growth low and dense; reverse olive or yellow. No germination at 58C and no growth at 378C. Conidiophores borne from aerial hyphae, characteristically of indeterminate form, sometimes terminating in well-defined penicilli with 2–4 metulae, sometimes bearing metulae in a random manner, with or without solitary phialides as well, less commonly giving the impression of monoverticillate penicilli borne perpendicular to fertile hyphae, but always with at least two terminal metulae; stipes smooth walled, of irregular and often indeterminate length; metulae usually long, terminating in well defined, thin walled vesicles; phialides ampulliform, 5–10 mm long; conidia ellipsoidal, 2.5–3.2 mm long, with surfaces finely to distinctly roughened, borne in long, irregular columns. Distinctive features. A well defined species, Penicillium fellutanum is distinguished microscopically by conidiophores with irregularly located metulae of variable length, terminating in definite vesicles, and macroscopically by closely textured colonies, dark green conidia and failure to grow at either 5 or 378C. Moreover growth on MEA is slower than on CYA and little faster than on G25N. Physiology. Penicillium fellutanum is a slow growing xerophile. Snow (1949) reported a minimum aw for germination of 0.80 at 258C and, in good agreement, Hocking and Pitt (1979) reported germination at 0.78 aw after 89 days. As noted above, P. fellutanum has a relatively narrow temperature range for growth, within the limits of 5 and 378C. Mycotoxins. Mycotoxins are not known to be produced. Ecology. Penicillium fellutanum has been isolated from a wide variety of dried foods, which appear to

212

7

Penicillium and Related Genera

Fig. 7.25 Penicillium fellutanum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

be a major habitat, including barley, wheat and flour, nuts, beans and sultanas. It has been isolated also from frozen fruit pastries and miso (see Pitt and Hocking, 1997). We isolated P. fellutanum at low levels from Thai cashews, and Indonesian maize, peanuts and milled rice (Pitt et al., 1993, 1998a). Additional species. Penicillium waksmanii K.M. Zalessky has much in common with P. fellutanum, including the variable and irregular penicillus structure and closely textured colonies. The principal

distinctions are that P. waksmanii often grows much faster on MEA (20–35 mm), germinates at 58C and produces greenish grey (25C2-26E3) spherical conidia. Mycotoxins are not produced. Habitats in foods appear to be similar to those of P. fellutanum, but P. waksmanii has been reported less frequently in the literature. Reports include ham during ripening (Spotti et al., 1989), peanuts (Pitt et al., 1993) and pistachios (Heperkan et al., 1994). References. Pitt (1979b, 2000).

7.7

Genus Penicillium

Penicillium janczewskii K.M. Zalessky Fig. 7.26 Penicillium nigricans Bainier

Colonies on CYA 25–32 mm diam, radially sulcate, deep, dense to moderately floccose; mycelium usually white, in some isolates pale yellow; conidial formation variable, in lightly sporing isolates coloured greenish grey (25B2-3), but in those sporing more heavily much darker, dull green (28E3); clear exudate and brown soluble pigment sometimes produced; reverse coloured brown, dark brown, orange or deep reddish orange. Colonies on MEA 18–24 mm diam, less commonly 30 mm, plane or radially sulcate, moderately deep to deep, dense to floccose; mycelium white to pale yellow, occasionally pale orange or pinkish; conidial production moderate to heavy, marginal areas sometimes bluish but greenish grey (25-26D-E2) elsewhere; orange or yellow soluble pigment sometimes produced; reverse typically Salmon (6A4-5) or, in the presence of soluble pigment, dark orange (5A8). Colonies on G25N 15–20 mm diam, occasionally only 12 mm, typically closely radially sulcate, velutinous to floccose; mycelium white to pale yellow; reverse usually pale yellow to dull brown. At 58C, typically germination by a proportion of conidia, less commonly general germination or microcolony formation. At 378C, typically no growth, occasionally colonies up to 10 mm diam formed. Conidiophores borne from aerial hyphae, stipes with thin smooth walls, commonly 50–200 mm long, but in some isolates much longer, and in the limit becoming indistinguishable from fertile hyphae bearing short conidiophores, characteristically bearing a terminal tetrad of divergent metulae, but frequently less regular in pattern, with intercalary rami and metulae commonly present, the latter appearing as short monoverticillate conidiophores; metulae 8–15 mm long, sometimes apically swollen; phialides ampulliform, 6–8 mm long, with short collula; conidia spherical, 2.5–3.5 mm diam, spinose, appearing olive brown, borne in short, poorly defined columns. Distinctive features. Despite considerable isolate to isolate variation, Penicillium janczewskii usually can be recognised by spherical spinose conidia, and metulae which are often apically swollen and characteristically occur in tetrads. However, in some isolates the most characteristic feature of the

213

conidiophores may appear to be a total lack of order in their structure. Colonies are floccose to a greater or lesser extent; conidial production is light to moderate; when moderate, conidial colour is dark green. Taxonomy. Raper and Thom (1949: 325) used the name Penicillium nigricans for this taxon. However, P. janczewskii is the correct name for this species (Pitt, 1979b; Pitt and Samson, 1993), and it is now in general use. Physiology. With an optimum temperature for growth near 258C, Penicillium janczewskii grows weakly at 58C and not above 338C (Domsch et al., 1980). This species is among the most xerophilic Penicillia, germinating down to 0.78 aw at 258C (Hocking and Pitt, 1979). Mycotoxins. Penicillium janczewskii has been reported to produce griseofulvin (El-Banna et al., 1987b; Nicoletti et al., 2007) and penitrem A (di Menna et al., 1986; Pitt and Leistner, 1991; Frisvad et al., 2006a). However, as this species is rarely if ever involved in food spoilage, significant mycotoxin production in the human food supply is unlikely. Ecology. All the evidence indicates that Penicillium janczewskii is a soil fungus, present in foods only as a contaminant, as we have no record of it having caused food spoilage. It has been reported from barley, wheat, flour, peanuts, pecans, pistachios, soybeans, dried beans and meats (see Pitt and Hocking, 1997). Additional species. Penicillium canescens Sopp is closely related to P. janczewskii, and in fact these species interface (Pitt, 1979b: 253). Both species grow at similar rates under the standard conditions and produce similar pigmentation, although P. canescens is often more strongly coloured. Microscopically, typical isolates are easily distinguished: P. canescens produces rough walled stipes and smooth conidia, while P. janczewskii has smooth stipes and rough walled conidia. Like P. janczewskii, P. canescens has been reported to produce griseofulvin (Nicoletti et al., 2007). This is a soil fungus, present in foods as a contaminant. P. canescens has been isolated from similar substrates to P. janczewskii, though less frequently, and also from rice, field peas (our data) and from cheese (Leistner and Pitt, 1977). References. Pitt (1979b, 2000); Domsch et al. (1980).

214

7

Penicillium and Related Genera

Fig. 7.26 Penicillium janczewskii (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

Penicillium janthinellum Biourge

Fig. 7.27

Colonies on CYA 35–50 mm diam, radially sulcate or irregularly wrinkled, floccose, mycelium dense, coloured white, greyish, buff, pale yellow or pale pink, and overlaid by conidiophores, varying from inconspicuous to abundant; conidial production very light to moderate, in the latter case coloured greyish green to dull green (25-27C-D3-4), sometimes appearing more yellow or olive because of

the coloured mycelium; limited amounts of clear to brown exudate and reddish brown soluble pigment sometimes produced; reverse colours variable, pale yellow or yellow brown to reddish brown, or occasionally brilliant dark green. Colonies on MEA usually 35–45 mm diam, floccose, lacking the density of colonies on CYA; mycelium white or buff; conidial colours similar to those on CYA; yellow or brown soluble pigment occasionally produced; reverse pale, brownish, deep brown, dark green or

7.7

Genus Penicillium

quite commonly pink, centrally or in sectors, occasionally even bright red. Colonies on G25N 10– 18 mm diam, typically plane, low or umbonate, velutinous to floccose; mycelium white or yellow; conidia grey green; reverse pale, yellow, brown or pinkish. Sometimes germination at 58C. At 378C, colonies 10–30 mm diam, dense and velutinous; clear exudate and brown or reddish soluble pigment sometimes produced; reverse pale, yellow, brown or reddish brown.

215

Conidiophores borne from surface or aerial hyphae, stipes smooth and slender, thin walled and easily bent, typically 200–400 mm long, but in some isolates also short, 30–70 mm, usually terminating in an irregular to regular verticil of 2–3 metulae, characteristically also with subterminal and intercalary metulae, the latter intergrading with short monoverticillate conidiophores, in some isolates long monoverticillate conidiophores present as well; phialides ampulliform, 7–11 mm long, typically

Fig. 7.27 Penicillium janthinellum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d, e, f) penicilli, bars = 10 mm; (g) conidia, bar = 5 mm

216

7

with long slender collula; conidia most often spherical, but sometimes short pyriform to ellipsoidal, 2.2–3.0 mm diam or long, with smooth to finely roughened walls, borne in short to moderately long disordered chains. Distinctive features. Penicillium janthinellum is surely the most difficult of all Penicillia to define. Colonies on CYA and MEA at 258C grow rapidly; growth typically occurs at 378C; colonies are usually floccose and conidia rather sparsely produced. Penicilli, although usually biverticillate, are so irregular as to often appear monoverticillate. Stipes are delicate and smooth walled. Phialides characteristically have long slender necks and bear smooth to finely roughened conidia. Physiology. Temperature requirements for this species are approximately known: minimum temperature for growth, above 58C, optimum 25–308C, good growth at 378C (Pitt, 1973). No water relations studies have been reported, but this species is probably not a xerophile. Mycotoxins. Penicillium janthinellum produces a series of tremorgenic toxins known as janthitrems (Gallagher et al., 1980). Closely related in chemical structure to the penitrems (de Jesus et al., 1984), janthitrems might be expected to show similar toxicities, however, toxicological data are not available. As P. janthinellum is uncommon in foods, these toxins are unlikely to be important in human health. Ecology. As an ubiquitous soil fungus, the presence of Penicillium janthinellum on foods is adventitious. Isolations have been made occasionally from a wide variety of foods: dates and almonds, maize, dried beans, peanuts, pistachios, barley, fermented and cured meats, biltong and frozen fruit pastries (see Pitt and Hocking, 1997). There appear to be no records of food spoilage by this species. References. Pitt (1979b, 2000); Domsch et al. (1980).

Penicillium oxalicum Currie & Thom Fig. 7.28 Colonies on CYA 35–60 mm diam, plane or radially sulcate, velutinous or lightly floccose in central areas; mycelium usually inconspicuous, in floccose areas white or pale yellow, but the underlying surface growth coloured Salmon (6A4); conidial production typically very abundant, appearing as a

Penicillium and Related Genera

continuous layer of long, closely packed chains under the low power microscope, and breaking off in masses if jarred, greyish green (25-26C3) at the margins, then dull green (27D3-28E4) or olive (1E3-4) towards the centres; exudate limited, clear, or absent; reverse pale to yellow, brown, orange or pinkish. Colonies on MEA variable in size, 20–50 mm diam, plane or lightly radially sulcate, strictly velutinous; conidia very abundant, forming readily detached masses; colours similar to those on CYA except reverse sometimes greenish. Colonies on G25N 12–16 mm diam, plane or wrinkled, velutinous; mycelium white or salmon; reverse pale, greenish, olive or salmon. At 58C, germination by a proportion of conidia, or no germination. At 378C, colonies 10– 40 mm diam, deeply radially sulcate and often centrally wrinkled, velutinous; mycelium white; reverse olive or brown. Conidiophores borne from surface mycelium, stipes mostly 200–400 mm long, with thin, smooth walls, characteristically terminating in verticils of 2–4 closely appressed metulae; metulae 15–25(–30) mm long; phialides acerose, 10–15(–20) mm long, with short collula; conidia ellipsoidal, very large, 3.5–5.0(–7) mm long, with walls smooth or rarely finely roughened, borne in long, closely packed columns. Distinctive features. Of all the cosmopolitan Penicillia, this species is perhaps the most obviously distinctive. Colonies usually grow rapidly on CYA at 25 and 378C, are strictly velutinous, and produce prodigious numbers of conidia. Under low magnifications, the conidia can be seen to lie in closely packed, readily fractured sheets, and to have a uniquely shiny, even silky, appearance. Under high magnification, the large penicilli and large smooth walled ellipsoidal conidia also are distinctive. Physiology. Mislivec and Tuite (1970b) reported 88C to be the minimum growth temperature for Penicillium oxalicum, with an optimum near 308C. Judged from rapid growth at 378C, the maximum temperature for growth is in excess of 408C. The minimum aw for germination has been reported as 0.86, both in glucose media at 23 and 308C (Mislivec and Tuite, 1970b) and in NaCl media at 258C (Hocking and Pitt, 1979). P. oxalicum was one of several Penicillium species associated with production of volatiles in mouldy wheat (Sinha et al., 1988). Mycotoxins. Secalonic acids are dimeric xanthones produced by a range of taxonomically

7.7

Genus Penicillium

217

Fig. 7.28 Penicillium oxalicum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

distant fungi (Cole and Cox, 1981; Frisvad and Thrane, 2004). Secalonic acid D is produced as a major metabolite of Penicillium oxalicum and is known to induce cleft palate in animals and potentially in humans (Dhulipala et al., 2005; Reddy, 2005). It has been found in nature, in grain dusts, at levels of up to 4.5 mg/kg (Ehrlich et al., 1982) and was also reported from grain dust collected in Belgian cereal storage facilities (Tangni and Pussemier, 2007). The possibility that such levels can be toxic to grain handlers cannot be ignored.

Ecology. As a result of its rapid growth at 378C, Penicillium oxalicum is widespread in tropical commodities and foods. We isolated it from Indonesian maize, red rice, paddy rice, cowpeas and sorghum, infecting 1% of all grains examined in each case (Pitt et al., 1998a) and at lower levels in peanuts, kemiri nuts, soybeans, mung beans, black pepper, coriander and milled rice in Thailand, Indonesia and/or the Philippines (Pitt et al., 1993, 1994, 1998a). A major niche for Penicillium oxalicum is preharvest maize (see Pitt and Hocking, 1997). Koehler

218

7

(1938) suggested that entry of this species to ripening maize ears was through insect damage or wounds. P. oxalicum has also been recorded in stored maize, though at lower incidence (Amusa et al., 2005; Askun, 2006). This species is a pathogen on yams (Okigbo, 2003) and greenhouse cucumbers (Menzies et al., 2005), is considered to be the dominant fungus on cassava (Adegoke et al., 1993) and is of common occurrence on copra (Zohri and Saber, 1993) and cashew nuts (Freire and Kozakiewicz, 2005). It has also been reported from a wide variety of other foods: barley, wheat, flour, pecans, hazelnuts, walnuts, peppercorns, soybeans, spices, fermented sausages and biltong (see Pitt and Hocking, 1997). The temperature and water relations of this species make it competitive with Aspergillus flavus except at very low aw, and it occupies a similar range of habitats. It is not such a common cause of spoilage, however. References. Pitt (1979b, 2000); Domsch et al. (1980).

Penicillium paxilli Bainier

Fig. 7.29

Colonies on CYA 30–35 mm diam, occasionally only 25 mm, radially sulcate, velutinous to lightly floccose; mycelium white at the margins, sometimes centrally greyish orange; conidial production moderate, at the margins greyish turquoise (24D3-4), centrally dull green (25-26E3-4); clear to red brown exudate and red brown soluble pigment often produced; reverse pale to brownish orange. Colonies on MEA usually 25–30 mm diam, occasionally less, plane to deeply radially sulcate, velutinous or occasionally floccose centrally; mycelium white to buff; conidial production moderate to heavy, coloured as on CYA; exudate and soluble pigment produced occasionally, red brown; reverse pale to brownish orange. Colonies on G25N 16–20 mm diam, radially sulcate, deep, dense to floccose: mycelium white; exudate and soluble pigment occasionally produced, red brown; reverse pale to brownish orange or reddish brown. At 58C usually no germination; less commonly germination or formation of microcolonies. At 378C, typically no growth, occasionally colonies up to 10 mm diam formed. Conidiophores borne from subsurface or surface hyphae, stipes usually 200–400 mm long but sometimes much longer, walls smooth to finely

Penicillium and Related Genera

roughened, typically bearing terminal verticils of 5–8 closely appressed metulae, but in some isolates subterminal metulae or even a subterminal ramus also occurring; metulae 10–15 mm long, sometimes enlarging at the apices; phialides ampulliform, 7–9 mm long, with short collula; conidia subspheroidal, 2.2–3.0 mm diam, smooth walled, borne in disordered chains. Distinctive features. Penicillium paxilli produces moderately sized, usually velutinous colonies, with terminal penicilli comprised of a crowded cluster of metulae. Stipes which are smooth or nearly so and the absence of sclerotia distinguish this species from P. raistrickii and P. novae-zeelandiae. Physiology. Little is known about the physiology of this species. It grows poorly, if at all, at 5 and 378C, and is probably not a xerophile. Mycotoxins. Penicillium paxilli has been reliably reported to produce two toxic compounds: verruculogen and paxilline. Verruculogen, a tremorgenic toxin with the same active indole moiety as penitrem A, but of quite different structure (Cole, 1981a), is almost equally toxic (Cole et al., 1972). However, the absence of reports of natural occurrence means it may be regarded as a mycotoxin of little consequence in foods and feeds, although not because it lacks toxicity. See P. simplicissimum for further details. Paxilline is also a tremorgen, but structurally quite different from the other tremorgens made by Penicillium species, and of lower toxicity. Again, this toxin has not been found from natural sources. Paxilline and verruculogen were assessed for their genotoxicity to human lymphocytes and in a mammalian microsome assay. Verruculogen tested positive in the mammalian microsome assay and paxilline caused DNA damage in human lymphocytes (Sabater-Vilar et al., 2003a). It is of interest that isolates of Penicillium paxilli producing verruculogen appear to come from Australia, while those producing paxilline are found in the United States (Cockrum et al., 1979). Ecology. Like most other species in subgenus Furcatum, Penicillium paxilli is a soil fungus and its occurrence in foods is usually adventitious. However, it has been reported to cause spoilage of garlic (Roy et al., 1977). Although there are few other references in the literature, it is our

7.7

Genus Penicillium

219

Fig. 7.29 Penicillium paxilli (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

experience that this species has a widespread though sparse distribution in foods. It has caused visual spoilage of cheese (our observations), and we isolated it at low levels from a wide range of Southeast Asian foods: peanuts from Thailand and dried fish, maize, paddy rice, peanuts, sorghum and soybeans from Indonesia (Wheeler et al., 1986; Pitt et al., 1993, 1998a). References. Pitt (1979b, 2000).

Penicillium raistrickii G. Sm.

Fig. 7.30

Colonies on CYA 35–45 mm diam, plane or nearly so, moderately dense to deeply floccose; mycelium usually white, sometimes centrally pale yellow (13A3); sclerotia usually abundant, enveloped by the mycelium; conidial formation often sparse and confined to central areas, sometimes moderate and then coloured dull green (26-27D3); clear exudate usually

220

present; reverse pale, yellow or yellow brown. Colonies on MEA usually 40–45 mm diam, occasionally as small as 30 mm, plane, sparse, velutinous to floccose; mycelium white to pastel yellow (3-4A3-4); sclerotia usually abundant, superficial or surrounded by wefts of mycelium; conidial production sparse, coloured as on CYA; exudate produced occasionally, red brown; reverse pale or yellow brown. Colonies on G25N 18–26 mm diam, plane or sulcate, often very deep and floccose at the centres; mycelium

7

Penicillium and Related Genera

white; reverse pale. At 58C, at least germination; usually colonies of 2–4 mm diam produced, of white mycelium. No growth at 378C. Sclerotia white or buff, rarely brown, 150–250 mm diam, texture firm to sclerotioid. Teleomorph not known. Conidiophores borne from subsurface mycelium and also from aerial hyphae adjacent to sclerotia, stipes usually 300–500(–800) mm long but from aerial hyphae sometimes much shorter, walls thick and roughened, bearing exclusively or

Fig. 7.30 Penicillium raistrickii (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) conidia, bar = 5 mm

7.7

Genus Penicillium

predominantly terminal biverticillate penicilli, in some isolates accompanied by short subterminal rami; terminal metulae in divergent verticils of 3– 5, apically inflated or clavate, of uniform appearance, 12–15 mm long, usually enlarging to 5–7 mm at the apices, sometimes rough walled; phialides acerose-ampulliform, 7–9 mm long, narrowing abruptly to short collula; conidia spherical to subspheroidal, 2.0–3.0 mm diam, smooth walled, borne in long, divergent columns. Distinctive features. Penicillium raistrickii grows rapidly and produces long, rough walled stipes and terminal biverticillate penicilli with apically inflated metulae. Almost all isolates produce abundant pale to brown sclerotia. Physiology. Little is known about the physiology of this species. Isolates are just capable of growth at 58C, but not at 378C. It is probably not a xerophile. Mycotoxins. Griseofulvin is a valuable antibiotic, but is appreciably toxic, and must also be classed as a mycotoxin. Penicillium raistrickii is one species producing this compound. This species may also make other unknown tremorgenic toxins (Patterson et al., 1979). Ecology. Basically a soil fungus, Penicillium raistrickii has a widespread though sparse distribution in foods. This species was isolated from contaminated capsicums (Martı´ n et al., 2005). We isolated it at low levels from Indonesian dried fish (Wheeler et al., 1986) and from Thai maize (Pitt et al., 1993). Other isolates in our culture collection have come from a mandarin fruit, from flour and from peanuts. Additional species. Perhaps a rarer species, Penicillium novae-zeelandiae J.F.H. Beyma appears to be very closely related to P. raistrickii. The major difference is that P. novae-zeelandiae characteristically produces irregular black sclerotia under the agar surface. What little is known of its physiology shows similarity to P. raistrickii. An isolate of P. novae-zeelandiae was found to produce an uncharacterised tremorgenic toxin (di Menna and Mantle, 1978). We have isolated P. novae-zeelandiae occasionally from a variety of foods, of note from ‘‘thread mould’’ spoilage of vacuum-packed cheese blocks (Hocking and Faedo, 1992), dried fruit and grape juice. References. Pitt (1979b, 2000).

221

Penicillium simplicissimum (Oudem.) Thom Fig. 7.31 Penicillium paraherquei S. Abe ex G. Sm. Penicillium piscarium Westling Penicillium pulvillorum Turfitt

Colonies on CYA typically 40–50 mm diam, radially sulcate; mycelium dense, white or occasionally buff; conidial production absent in some isolates, light to moderate in others, typically coloured greyish green to dull green (25C-D4) but in some isolates centrally or predominantly yellowish grey (2-3C2); sometimes clear exudate and occasionally red brown soluble pigment present; reverse typically pale, but sometimes in greyish yellow shades such as chamois or khaki (4B-D5). Colonies on MEA typically 40–50 mm diam, plane, up to 2–3 mm deep, velutinous to floccose; mycelium white; conidial production moderate to heavy, greyish green to dull green (26-29C-E3-4); reverse pale or sometimes yellow brown. Colonies on G25N 12–18 mm diam, plane, sulcate or wrinkled, velutinous to floccose; mycelium white; reverse pale, yellowish or olive. At 58C, no germination to germination, occasional isolates forming microcolonies. At 378C, colonies usually 20–30 mm diam, occasional isolates weaker, 5–10 mm or even absent, sulcate to deeply wrinkled, of white to pinkish mycelium, rarely with grey or greenish grey conidia present; reverse pale or brownish. Sclerotia occasionally produced on CYA or MEA at 258C, spheroidal or irregular, commonly 200–500 mm diam, brown, of soft texture. Teleomorph not known. Conidiophores borne from surface or aerial hyphae, stipes commonly 400–800 mm long, usually rough walled, bearing regular to irregular verticils of metulae and occasionally a well defined ramus, but only rarely short conidiophores or metulae in other than terminal or subterminal positions; metulae in verticils of 2–5, 12–20 mm or more long, rough walled; phialides ampulliform, mostly 7–9 mm long, narrowing abruptly to long collula; conidia variable, commonly ellipsoidal, but at times spherical, subspheroidal or pyriform, 2.5–4.0 mm long, with walls roughened or spinulose, borne in disordered chains.

222

7

Penicillium and Related Genera

Fig. 7.31 Penicillium simplicissimum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

Distinctive features. Isolates of Penicillium simplicissimum grow rapidly at 258C, forming comparatively deep colonies due to long conidiophores. The conidiophores are characteristic: stipes are long and coarse with rough walls and bear terminal or subterminal metulae and occasionally rami. Short conidiophores or intercalary metulae are uncommon. Taxonomy. Controversy over the correct name and circumscription of this species (Stolk and

Samson, 1983; Pitt, 2000; Frisvad et al., 1990b) has been resolved by maintaining the neotypification by Pitt (1979b), based on a herbarium specimen of Jensen’s from Cornell University (Pitt and Samson, 1993). A recent taxonomic study of Penicillium simplicissimum and related species (Tuthill et al., 2001) concluded that P. simplicissimum encompasses several distinct species and that the name should be applied more narrowly within this group. However,

7.7

Genus Penicillium

as all of those species are very similar, and it is not known which occur in foods, the concept of P. simplicissimum used here is the same as that of Pitt (1979b, 2000). Physiology. Isolates of Penicillium simplicissimum from different parts of the world show variation in growth at higher temperatures. Tropical isolates grow strongly at 378C; some from cooler regions of Australia grow at most weakly at this temperature. This probably reflects the report that P. simplicissimum actually includes several species (Tuthill et al., 2001). This species is not a xerophile, germinating down to 0.86 aw only (Hocking and Pitt, 1979). Mycotoxins. Penicillium simplicissimum has been reported to produce the mycotoxins verruculogen (Pitt, 1979c), fumitremorgen B (El-Banna et al., 1987b), penicillic acid and viridicatumtoxin (El-Banna et al., 1987b; Pitt and Leistner, 1991). Verruculogen, of quite different structure from the penitrems but containing the same active indole moiety (Cole, 1981b), is almost equally toxic (Cole et al., 1972, reported as P. verruculosum). Fumitremorgen B is closely related to verruculogen and can be expected to have a similar toxicity. However, natural production of these toxins by this species has not been shown, and their significance in human or animal health is doubtful. The importance of viridicatumtoxin is unknown (see P. aethiopicum). In splitting Penicillium simplicissimum into several species, Tuthill et al. (2001) showed that fumitremorgens, verruculogen and viridicatumtoxin are produced by P. brasilianum, not P. simplicissimum. However, it is not yet clear which of these species occurs in foods. This species appears to be resistant to formaldehyde. Isolates in our collection and at the International Mycological Institute, Egham, Surrey, UK, have come from 10% formalin solution or preserved specimens. Several IMI isolates have also come from polyvinyl chloride or polyester plastics. Ecology. Penicillium simplicissimum appears to have its primary habitat in decaying vegetation, and perhaps soil. Its occurrence on foods is therefore not as a spoilage fungus, but as a contaminant. It has been reported from country cured hams (Monte et al., 1986), cashews, peanuts and mung beans (Pitt et al., 1993, 1994). Isolates in our collection have also come from spring water, sorghum malt and fermented sausage. References. Pitt (1979b,c, 2000).

223

7.7.3 Penicillium subgenus Penicillium (Penicillium Sect. Asymmetrica Raper and Thom)

In Penicillium subgenus Penicillium, penicilli are predominantly terminal terverticillate structures, i.e. phialides are always borne on metulae and the metulae usually are borne on well -defined terminal rami. Biverticillate and quaterverticillate penicilli are also produced by some species. Phialides are typically ampulliform (flask shaped), but are cylindroidal to acerose (needle shaped) in a few species. Colonies on G25N normally exceed 18 mm diam, although they are consistently less in one species treated here, P. digitatum. Only P. aethiopicum and P. chrysogenum sometimes grow at 378C. Growth at 58C is always positive, and usually strong: most species produce macroscopic colonies in 7 days. By far the most important subgenus of Penicillium with respect to food spoilage, subgenus Penicillium is also by far the most difficult taxonomically, both because there are numerous species and because apparent differences between species are small. For a number of years, the taxonomy of this subgenus remained controversial, with the systems of Raper and Thom (1949), Samson et al. (1976) and Pitt (1979b) all being commonly used. During the 1980s, students of Penicillium sought consensus on the classification of this subgenus. Secondary metabolite profiles (Frisvad, 1981; Frisvad and Filtenborg, 1983) and isoenzyme patterns (Cruickshank and Pitt, 1987) provided independent evidence about species concepts. Integration of classical morphology with these new approaches led to consensus for most species (Samson and Pitt, 1985). Later, a list of ‘‘Names in Current Use’’ (NCU) was established for Penicillium species (Pitt and Samson, 1993; Pitt et al., 2000). This now provides much needed stability. All names used here for species in this subgenus are protected by this NCU. Recently, Samson and Frisvad (2004) have published a comprehensive revision of this subgenus and increased the number of included species from 23 to 58. Where these additions affect species included here they are discussed under the relevant species. Identifications. It is usually relatively easy to decide whether an isolate belongs in subgenus Penicillium, but accurate identifications to species level require care and, often, acquired skill. In the

224

keys used here, the roughness or smoothness of stipes and small differences in colour will affect the disposition of an isolate. Perhaps the most important point to note in the keys which follow is that species which have perfectly smooth stipe walls under the light microscope at 400–600 magnification are distinguished from those in which some stipes, though not necessarily all, are finely to distinctly roughened. Roughening of stipe walls is often more readily seen in wet mounts made from cultures grown on MEA. A second point to note is that judgment of colour should be made in daylight or artificial daylight conditions, i.e. in the latter case, under daylight type fluorescent tubes, not incandescent lamps. The use of the Methuen ‘‘Handbook of Colour’’ (Kornerup and Wanscher, 1978) will greatly assist colour differentiation of species in this subgenus. Some of the species accepted here are undoubtedly closely related and intermediate isolates will be encountered from time to time. Nevertheless, with care it should be possible to key out and recognise the majority of isolates from foods. Neutral creatine sucrose agar. Creatine sucrose agar (CREA) was introduced as a new approach to the problem of identifying species in subgenus Penicillium (Frisvad, 1981, 1985). He pointed out that species in this subgenus fall into two groups: those with an affinity for proteinaceous foods and those which will grow vigorously on foods rich in carbohydrate. CREA was designed to permit differentiation between these two groups. It contained the organic base creatine as a sole source of nitrogen, sucrose as a source of carbon, and an indicator, bromocresol purple, to detect pH changes related to differential acid production (from sucrose) or base production (by liberation of ammonium ions from creatine). The concept was innovative and sound, but CREA was not easy to use in practice. A number of variations were introduced by Frisvad (1993), but without firm recommendations. A medium based on CREA, but of initial pH neutral to bromocresol purple and with a change in sucrose to creatine ratio to provide more positive responses by some species, was developed by Pitt (1993a) and termed Neutral Creatine Sucrose agar (CSN). The formula is given in Appendix.

7

Penicillium and Related Genera

CSN has proved to be very useful in identification of species in Penicillium subgenus Penicillium, as eight different types of responses have been observed among common species (Table 7.2; modified from Pitt, 1993a). Reaction to CSN has been included in the descriptions and keys which follow. If only occasional isolates from this subgenus are seen, use of CSN is not essential. However, for the laboratory which frequently isolates species from this subgenus, CSN can be of great assistance. CSN is of little value outside this subgenus. Use of Ehrlich reagent. In the quest for more rapid and reliable methods for identifying species in subgenus Penicillium, Lund (1995a) developed a simple test to distinguish among several species common in cheese and bread factories in Denmark. The test used Ehrlich reagent to visualise production of indole compounds as secondary metabolites. Ehrlich reagent is made by dissolving 4-dimethylaminobenzaldehyde (2 g) in 96% ethanol (85 ml) and adding 10 N HCl (15 ml). To carry out the test, a cork borer (4 mm diameter) is used to cut an agar plug from a culture on CYA and then the plug is placed, mycelium side up, on a Petri dish lid. A strip (12  6 mm) of Whatman No. 1 filter paper is dipped in the Ehrlich reagent and then placed across the agar plug ‘‘like a propeller’’, with the agar plug in the centre of the strip, so the strip does not touch the Petri dish lid. The arrangement is then placed in an air stream in a fume hood and allowed to dry. A positive response is indicated by a violet ring in the filter paper, appearance after 2–6 min being a strong response, and after 7–10 min a weak one. Observation must be made within 10 min, as the ring fades after that time. Most isolates produce the colour on CYA after incubation for 7 days, however, a few require 14 days incubation. The test is of more value when positive than negative, therefore. Species for which this test is positive include Penicillium camemberti, P. commune, P. expansum and P. roqueforti (Lund, 1995a). A few other species produce yellow colours with this test, including P. crustosum (Lund, 1995a). Physiology. As noted above, species in subgenus Penicillium are very important in food spoilage. They are able to grow at low temperatures and quite low water activities and are of universal occurrence in cereals, refrigerated foods and

7.7

Genus Penicillium

225

Table 7.2 Responses of common species of Penicillium subgenus Penicillium to Neutral Creatine Sucrose agar (CSN) after incubation for 7 days at 258Ca Species

Colony growth

Colony diam (mm)

Penicillium aethiopicum P. allii P. aurantiogriseum P. brevicompactum

Moderate

15–20

Weak Moderate Weak

Medium reaction

Colony reverse

Acid + brown

15–20 15–25 8–14(–20)

Acid, margins sometimes neutral Acid Acid + brown Neutral to weakly acid

15–20 12–18

Alkaline, margins neutral Neutral or weakly acid

P. commune

Strong Moderate to strong Strong

20–26

P. crustosum

Strong

25–30

P. digitatum P. echinulatum P. expansum

Weak Strong Strong

4–10 22–25 24–30

Alkaline, sometimes weakly acid Acid, occasionally alkaline Neutral Alkaline Acid

P. glandicola P. griseofulvum P. hirsutum P. hordei P. italicum P. nalgiovense P. olsonii P. roqueforti

Moderate Weak Strong Strong Weak Weak Weak Strong

12–18 18–24 24–30 20–24 10–20 10–18 6–14 25–40

Weakly acid to acid Neutral Acid Acid Neutral Neutral to weakly acid Neutral Variable

P. solitum P. ulaiense P. verrucosum P. viridicatum

Strong Weak Weak Moderate to strong

18–22 4–8 10–15 15–22

Usually alkaline Neutral Neutral Acid

P. camemberti P. chrysogenum

Acid Acid + brown Neutral (occasionally weakly acid) Alkaline Neutral to weakly acid Alkaline Alkaline, occasionally weak Neutral Alkaline Acid + brown, occasionally alkaline Acid + brown Neutral Acid (+ brown) Acid + brown Neutral Neutral to weakly acid Neutral Alkaline, occasionally neutral Alkaline Neutral Neutral Acid (+ brown)

a

Acid, yellow; weakly acid, pale yellow; neutral, grey, sometimes with a reddish cast; weakly alkaline, grey violet; alkaline, violet or reddish violet. Brown responses are from naturally produced pigments.

many other environments. Controlling these fungi in many types of bulk stored food commodities relies on a combination of low water activity and low temperature: even marginal errors in these controls may sometimes lead to high losses. Mycotoxins. With the possible exception of the fruit-rotting species Penicillium digitatum, P. italicum, P. solitum and P. ulaiense, all of the species under consideration here produce mycotoxins. Moreover, mycotoxin production appears to occur more consistently than in most other genera: a large majority of the isolates encountered from subgenus Penicillium will be mycotoxigenic. Fortunately, most of the toxins are believed to have relatively low potency or are not produced under conditions occurring in foods.

The question of species–mycotoxin associations for species in Penicillium subgenus Penicillium, considered a major problem when the first edition of this book was published, has been clarified. A comprehensive collaborative study, in which more than 1500 Penicillium isolates were critically examined both taxonomically and for mycotoxin production, greatly assisted this process (El-Banna et al., 1987b; Pitt and Leistner, 1991). When combined with data from Frisvad and Filtenborg (1989) and Samson and Frisvad (2004), this information has enabled recognition of the fact that species–mycotoxin associations in this genus are mostly quite specific. These data have been incorporated below.

226

7

Penicillium and Related Genera

Key to Penicillium subgenus Penicillium species included here 1

Conidia white or pale grey green Conidia more strongly coloured, blue, green or grey

2 (1)

Stipes often rough walled, conidia up to 5 mm diam; on CSN, reaction alkaline Stipes consistently smooth walled, conidia not usually exceeding 3.5 mm diam; on CSN, reaction neutral to weakly acid

3 (1)

4 (3)

Stipes on CYA and MEA smooth walled, or at most very finely roughened Stipes commonly rough walled, especially on MEA Conidia borne as cylinders, with at least a proportion remaining so at maturity Conidia borne as ellipsoids or spheres and remaining so at maturity

5 (4)

Conidia olive, longer than 6 mm Conidia green, shorter than 6 mm

6 (5)

Colonies on CYA and MEA exceeding 30 mm diam Colonies on CYA and MEA not exceeding 30 mm diam

7 (4)

Penicilli with 3 or more rami in large, compact terminal penicilli Penicilli with 1–2 rami, penicillus structure variable

8 (7)

Penicilli complex or irregular, reverse on MEA strongly coloured, deep orange brown to brown, phialides with short, broad collula; on CSN, reaction neutral to weakly acid Penicilli well defined, mostly terverticillate, reverse on MEA pale or dull yellow or brown, phialides with relatively long, narrow, collula; CSN response variable

2 3 P. camemberti P. nalgiovense 4 16 5 7 P. digitatum 6 P. italicum P. ulaiense (see P. italicum) P. olsonii 8

P. nalgiovense

9

9 (8)

Colonies on CYA exceeding 30 mm diam Colonies on CYA not exceeding 30 mm diam

10 14

10 (9)

Conidia grey blue Conidia grey green to green

11 12

11(10)

Colonies on CYA 35–45 mm diam, yellow exudate and soluble pigment often present, conidia ellipsoidal to subspheroidal; on CSN, medium response neutral to weakly acid Colonies on CYA 30–38 mm diam, exudate clear to pale brown, soluble pigment sometimes present, brown to reddish brown, conidia spherical to subspheroidal; on CSN, medium response acid, reverse acid plus brown

12(10)

13(12)

14 (9)

Reverse on CYA pale to deep brown; colonies on CSN usually exceeding 24 mm diam Reverse on CYA pale, yellow or orange; colonies on CSN less than 24 mm diam Colonies dull green, conidia broadly ellipsoidal, reverse on CYA bright golden yellow; on CSN reverse acid plus brown Colonies dark green, conidia mostly spherical to subspheroidal, reverse on CYA usually pale, sometimes orange or pale yellow; CSN reverse alkaline Phialides commonly 4.5–6 mm long; on CSN, response neutral Phialides exceeding 6 mm long; CSN response variable

P. chrysogenum

P. aurantiogriseum P. expansum 13 P. aethiopicum P. solitum P. griseofulvum 15

7.7

Genus Penicillium

15(14)

227

On CYA colonies dull green, penicilli very broad, metulae often apically inflated, conidia ellipsoidal; on CSN, reverse neutral to weakly acid On CYA, colonies dark green, penicilli not very broad, metulae not usually apically inflated, conidia mostly spherical to subspheroidal; on CSN, reverse alkaline

P. brevicompactum

P. solitum

16 (3)

Colonies on CYA exceeding 30 mm diam Colonies on CYA not exceeding 30 mm diam

17 24

17(16)

Conidia with walls finely roughened to rough or spinose Conidia smooth walled

18 19

18(17)

Mycelium inconspicuous, white, conidia dark green; growth on CSN strong, medium and reverse alkaline Mycelium yellow, conidia dull green; on CSN, medium and reverse acid

19(17)

20(19)

21(20)

22(19)

23(22)

24(16)

Growth on CSN strong, colonies more than 20 mm diam, medium and reverse alkaline Growth on CSN variable, medium and reverse acid, reverse sometimes also brown Colonies on CYA and MEA exceeding 40 mm diam, reverse on CYA and/or MEA often green to deep blue green; conidia up to 6 mm diam Colonies on CYA or MEA less than 40 mm diam, reverse pale or yellow to orange brown, conidia mostly 4 mm or less diam Colonies on CYA exceeding 35 mm diam and on MEA usually exceeding 30 mm diam; colonies on MEA often shedding masses of conidia when jarred Colonies on CYA and MEA not exceeding 35 and 30 mm diam, respectively; conidia adhering to colonies on MEA Colonies on CYA often exceeding 35 mm diam Colonies on CYA not exceeding 35 mm diam Exudate on CYA near maroon; mycelium often yellow; penicilli sometimes quaterverticillate Exudate on CYA pale yellow; mycelium white, penicilli terverticillate or somewhat irregular Conidia mostly ellipsoidal, dark green

Colonies on CYA and MEA exceeding 25 mm diam Colonies on CYA and MEA not exceeding 25 mm diam

Penicillium aethiopicum Frisvad

Fig. 7.32

Colonies on CYA 32–40 mm diam, finely radially sulcate, deeply fasciculate; mycelium white, visible only at the margins; conidial production moderate to heavy, dull green (25-26E-F4); copious exudate usually present, clear to pale brown; reverse closely sulcate, golden yellow near mustard yellow or maize

20 22

P. roqueforti

21

P. crustosum P. commune

23 P. viridicatum

P. hirsutum

P. allii (see P. hirsutum) P. glandicola (see P. brevicompactum) 25

Conidia near spherical, yellow green 25 (24)

P. echinulatum (see P. crustosum) P. hordei (see P. hirsutum)

P. viridicatum P. verrucosum

yellow (3-4B-C5-6). Colonies on MEA 32–40 mm diam, plane, low and dense, surface velutinous to granular; mycelium inconspicuous, white; conidial production moderate, dull green (26D-E4); reverse uncoloured to dull yellow brown. Colonies on G25N 22–24 mm diam, plane or sulcate, low to moderately deep, dense, usually fasciculate; mycelium white; sporulation moderate, dull green; soluble pigment

228

7

Penicillium and Related Genera

Fig. 7.32 Penicillium aethiopicum (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) conidia, bar = 5 mm

sometimes produced, pale orange (5A3); reverse pale, bright yellow or pale orange. At 58C, germination to colonies 3 mm diam. At 378C, no growth to colonies 10 mm diam, of dense white mycelium. On CSN, colonies 15–20 mm diam, growth moderate, medium reaction acid (pale yellow), sometimes neutral at the margins; reverse acid (yellow), plus brown. Conidiophores borne singly or more commonly in fascicles, mostly from subsurface hyphae, stipes 200–400

mm long, or of indeterminate length in fascicles, with smooth or at most very finely roughened walls, typically bearing appressed, terminal terverticillate penicilli; phialides ampulliform, 7–9 mm long, with short collula; conidia broadly ellipsoidal, 3.2–3.8 mm long, smooth walled, borne in long, sometimes well defined, columns. Distinctive features. Penicillium aethiopicum resembles P. chrysogenum in growth rates, including the ability to grow weakly at 378C; P. expansum, in

7.7

Genus Penicillium

229

producing fasciculate colonies; and both species, in producing conidiophores with smooth-walled stipes. The most obvious feature distinguishing P. aethiopicum from these other species is the formation of a closely sulcate golden yellow reverse on CYA in the absence of yellow soluble pigment. Penicilli are more compact and robust than those of P. chrysogenum, while brown pigmentation is much less than occurs in colonies of P. expansum on CYA. Growth on CSN resembles that of P. chrysogenum. Physiology. No studies on physiology are known to us. The ability to grow at 378C suggests a physiology similar to Penicillium chrysogenum. Mycotoxins. Penicillium aethiopicum produces griseofulvin (see P. griseofulvum) and viridicatumtoxin (Frisvad and Filtenborg, 1989). The latter is a moderately toxic compound, with an LD50 of 70 mg/kg when injected into mice. Its oral toxicity is unknown and its practical significance unclear. Ecology. Penicillium aethiopicum was originally described from barley from Ethiopia (Frisvad and Filtenborg, 1989). We found this species frequently in Southeast Asian commodities: in 16% of Indonesian kemiri nut samples, with a 4% infection rate in all nuts examined, in 18% of Indonesian cow pea samples, 9% of Indonesian peanut samples and 4% of Philippine maize samples. Incidence in some individual samples was high, up to 30–40% of all particles examined, providing a 1% infection level overall in each of these commodities. It was also present, at low levels, in Thai peanuts and cashews, Philippine soybeans and mung beans, and Indonesian milled rice, soybeans and mung beans (Pitt et al., 1993, 1994, 1998a). References. Frisvad and Filtenborg (1989); Samson and Frisvad (2004).

Penicillium aurantiogriseum Dierckx Fig. 7.33 Penicillium puberulum Bainier Penicillium cyclopium Westling Penicillium martensii Biourge Penicillium aurantiovirens Biourge Penicillium verrucosum var. cyclopium (Westling) Samson et al.

Colonies on CYA 30–37 mm diam, radially sulcate, moderately deep, texture smooth to granular; mycelium white, usually inconspicuous; conidial production moderate to heavy, greyish turquoise to dull green (24-25D-E3-4); exudate usually conspicuous, clear or pale brown; soluble pigment produced by

some isolates, brown to reddish brown; reverse pale, light to brilliant orange, or reddish to violet brown. Colonies on MEA 24–35 mm diam, plane or rarely radially sulcate, low and relatively sparse, surface texture finely granular; mycelium usually subsurface, occasionally conspicuous and then bright yellow; conidial production usually moderate to heavy, greyish turquoise to dull green (24-25D-E4-5); soluble pigment sometimes produced, yellow brown to reddish brown; reverse pale, orange, or reddish brown. Colonies on G25N 18–24 mm diam, usually radially sulcate, moderately deep, dense, texture granular; reverse pale, yellow or brown. At 58C, colonies 2–5 mm diam, of white mycelium. No growth at 378C. On CSN, colonies usually 15–25 mm diam, with moderately dense to dense growth, medium reaction acid (yellow); reverse acid (yellow) plus brown soluble pigment. Conidiophores borne singly or in fascicles, mostly from subsurface hyphae, stipes 200–400 mm long, or of indeterminate length in fascicles, with walls smooth to finely roughened, bearing terminal terverticillate or less commonly biverticillate penicilli; phialides slender, ampulliform, mostly 7–10 mm long; conidia spherical to subspheroidal, less commonly ellipsoidal, 3.0–4.0 mm long, with smooth walls, mostly borne in long, well defined columns. Distinctive features. Penicillium aurantiogriseum produces blue grey conidia on both CYA and MEA. Colonies on CYA grow relatively slowly (37 mm diam at most) and are dense and of granular texture. Colonies on MEA are relatively low and sparse, with distinctly granular margins. Stipes are usually smooth or nearly so. The response on CSN is distinctive: quite rapid growth, an acid reaction in medium and colony reverse, characteristically with brown colours also in the medium and beneath the colony. Taxonomy. Penicillium aurantiogriseum as described here is a relatively broad species. It has essentially the same circumscription as given by Pitt et al. (1986) and Pitt (2000), but narrower than that described by Samson et al. (1976) or Pitt (1979b). More recently, several other species were segregated from P. aurantiogriseum on the basis of secondary metabolite, morphological and ecological characters (Lund and Frisvad, 1994; Samson and Frisvad, 2004). We have maintained the concept of Penicillium aurantiogriseum described by Pitt et al. (1986) and Pitt (2000) as we believe that, with some practice, it

230

7

Penicillium and Related Genera

Fig. 7.33 Penicillium aurantiogriseum (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) conidia, bar = 5 mm

can be distinguished purely on morphological criteria from other species described in this book. If further differentiation is required, see Lund and Frisvad (1994) or Samson and Frisvad (2004). Physiology. Penicillium aurantiogriseum has a minimum temperature for growth near 28C, an optimum near 238C and a maximum near 308C (Armolik and Dickson, 1956; Mislivec and Tuite,

1970b). The minimum aw for growth is 0.81 (Mislivec and Tuite, 1970b). Growth at reduced aw is little affected by pH or substrate (Hocking and Pitt, 1979). Growth was stimulated by carbon dioxide levels above 10% (Magan and Lacey, 1984b). Growth was observed in 30 and 50% CO2 but not in atmospheres containing 70% CO2 (Zardetto, 2005).

7.7

Genus Penicillium

Considerable work has been carried out on the production of volatiles in stored grains by Penicillium aurantiogriseum. Major compounds reported include 1-propanol, 2-methyl-1-propanol, 3-methyl-1-butanol, 1-octen-3-ol and 3-octanone on barley, wheat and synthetic media (Sinha et al., 1988; Wilkins and Scholl, 1989; Borjesson et al., 1990; Larsen and ¨ Frisvad, 1995). The lipolytic activity of Penicillium aurantiogriseum was high, as judged by action on a medium containing 1% rapeseed oil (Magan et al., 1993). Lipase production was enhanced when a medium containing 0.5% yeast extract and 1% olive oil was supplemented with ammonium sulphate (Lima et al., 2003). Mycotoxins. Isolates of Penicillium aurantiogriseum commonly produce penicillic acid, verrucosidin and nephrotoxic glyopeptides (Frisvad and Samson, 2004). Accumulation of penicillic acid by P. aurantiogriseum is favoured by low temperatures, in the range 1–108C; production is faster at higher temperatures, but so is degradation (Ciegler and Kurtzman, 1970). Reports that Penicillium aurantiogriseum isolates may also produce roquefortine C, xanthomegnin or viomellein (El-Banna et al., 1987b; Frisvad and Filtenborg, 1989) are now believed to be incorrect (Samson and Frisvad, 2004) Ecology. Along with some other species in this subgenus, Penicillium aurantiogriseum is among the most commonly encountered fungi on earth. It is ubiquitous in maturing or drying crops, especially cereals and cereal products (see Pitt and Hocking, 1997). This association with cereals is very important: reports of P. aurantiogriseum from proteinaceous foods including meat and cheese are almost always in error (Frisvad and Filtenborg, 1989; Lund et al., 1995). It has also been isolated frequently from nuts (see Pitt and Hocking, 1997). Penicillium aurantiogriseum has been reported to cause spoilage of a variety of stored fruits and vegetables, including apples, pears, strawberries, grapes, melons, tomatoes, cassava and potatoes (see Pitt and Hocking, 1997). Not all such reports have been authenticated: it is likely that related species are responsible for some of these cases. Other reported sources include cold stored eggs, frozen fruit pastries, spices, dried beans and peas, soybeans, dried fruit, health foods (see Pitt and Hocking, 1997) and luncheon meats (Ismail and Zaky, 1999).

231

We isolated Penicillium aurantiogriseum from a variety of Southeast Asian food commodities, including maize, peanuts, cashews, soybeans, mung beans and rice, but only at low levels (Pitt et al., 1993, 1994, 1998a). References. Pitt (1979b, 2000); Samson et al. (1976) and Domsch et al. (1980), both under the name Penicillium verrucosum var. cyclopium; Samson et al. (1995); Samson and Frisvad (2004).

Penicillium brevicompactum Dierckx Fig. 7.34 Penicillium stoloniferum Thom

Colonies on CYA 20–30 mm diam, radially sulcate, moderately deep, dense, texture typically velutinous; mycelium white; conidial formation light to moderate, dull green (25-28D-E3); exudate usually present in minute droplets, often deeply embedded, but sometimes copious, pale to deep reddish brown; soluble pigment usually produced, reddish brown; reverse sometimes pale but more usually yellowish to reddish brown. Colonies on MEA 12–22 mm diam, plane or less commonly radially sulcate, usually velutinous; mycelium white; conidial production moderate to heavy, dull green to dark green (27-29E-F4), rarely paler or more bluish; exudate occasionally present, clear to reddish brown; reverse pale or brown. Colonies on G25N 14–22 mm diam, plane or radially sulcate, texture granular; clear exudate and red brown soluble pigment sometimes produced; reverse pale, yellow or reddish brown. At 58C, microcolonies to colonies up to 4 mm diam produced. No growth at 378C. Colonies on CSN 8–14 mm diam, occasionally larger, growth weak and sparse, medium response neutral to weakly acid (yellow); reverse neutral, occasionally weakly acid. Conidiophores borne from surface mycelium, stipes usually broad, 500–800 mm long, smooth walled, characteristically bearing compact, broad terverticillate penicilli, usually less than 40 mm long and 40–50 mm across the phialide tips, with quaterverticillate and biverticillate penicilli usually evident also; rami short and broad, often bent away from the axis; metulae in divergent clusters, short and broad, 9–15 mm long, typically apically inflated; phialides in divergent verticils, ampulliform, 6–9 mm long; conidia ellipsoidal, 2.5–3.5 mm long, with walls smooth to very finely roughened, borne in divergent and disordered chains.

232

7

Penicillium and Related Genera

Fig. 7.34 Penicillium brevicompactum (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) conidia, bar = 5 mm

Distinctive features. Penicillium brevicompactum produces compact (though not small) penicilli, often as wide as long. Metulae are short and broad, often apically inflated, fanning out so that the outermost phialides may point in almost diametrically opposed directions. As a result, conidial chains are borne over almost a hemisphere, in a shape suggesting Aspergillus. Physiology. The minimum and maximum temperatures for growth of Penicillium brevicompactum

are –2 and 308C, respectively (Mislivec and Tuite, 1970b), with an optimum near 238C. The minimum aw for germination and growth is 0.78 at 258C (Hocking and Pitt, 1979), categorising P. brevicompactum as one of the most xerophilic Penicillia. Mycotoxins. This species produces mycophenolic acid (Frisvad and Filtenborg, 1989; Frisvad and Samson, 2004), a weakly toxic compound with an oral LD50 of 700 mg/kg in rats (Cole and Cox, 1981). It is now used as an immunosuppressant in

7.7

Genus Penicillium

heart and kidney transplant patients (Bentley, 2000). The presence of mycophenolic acid in ginger due to infection by P. brevicompactum has been reported (Overy and Frisvad, 2005) but is unlikely to be of practical concern. Ecology. Although less common than some other species in this subgenus, Penicillium brevicompactum is nevertheless of widespread occurrence especially, because of its xerophilic nature, in dried foods: beans, soybeans, pecans, pistachios and peanuts, health foods and peppercorns (see Pitt and Hocking, 1997). It has also been isolated from Brazilian cashew nuts (Freire et al., 1999), Brazil nuts and black and white pepper (Freire et al., 2000). It commonly occurs in European meat products, hams, biltong (see Pitt and Hocking, 1997) and salami (Cantoni et al., 2007). P. brevicompactum can also spoil refrigerated products, such as cheese (Kure and Skaar, 2000; Kure et al., 2001; Hayaloglu and Kirbag, 2007). It has also been reported in tap water in Portugal (Gonc¸alves et al., 2006). In our laboratory P. brevicompactum has been isolated from a range of substrates including spoiled margarine, dairy products, fruit pure´e, curry paste, sumac, bakery products and bottled water. Penicillium brevicompactum can also behave as a weak pathogen having caused spoilage of stored apples, mushrooms, cassava, potato, pumpkin (see Pitt and Hocking, 1997), grapes (Bau et al., 2005, 2006; Patin˜o et al., 2007) yams (Aboagye-Nuamah et al., 2005) and lychees (our observations). This species has recently been isolated from fresh ginger, where it produced mycophenolic acid (Overy and Frisvad, 2005). This species was present at low levels in a variety of Southeast Asian commodities: rice, mung beans, soybeans, maize and peanuts (Pitt et al., 1993, 1994, 1998a). Additional species. Penicillium glandicola (Oudem.) Seifert and Samson (synonym P. granulatum Bainier) grows at similar rates to P. brevicompactum under all standard conditions. P. glandicola is distinguished by a granular texture on CYA and MEA, with small coremia apparent at the margins; stipes, rami and metulae with rough walls; and dark green conidia. On CSN, colonies are 12–18 mm diam, and growth is moderate, with an acid (yellow) reaction in both medium and colony reverse; brown soluble pigment is usually also produced.

233

Hocking and Pitt (1979) reported 0.86 as the minimum aw for germination and growth of Penicillium glandicola. This species has been reported to be one cause of taint in wine corks (Daly et al., 1984) and as one species producing 2,4,6 trichloroanisole, a cause of off flavour in coffee (Liardon et al., 1992). Penicillium glandicola is one potential source of penitrem A, patulin and roquefortine C, which may be produced in silage (Frisvad and Samson, 2004). Natural occurrence of these toxins in foods due to this species has not been reported. This is not a commonly occurring fungus, but it has been isolated from cereals sufficiently frequently to warrant mention here: from wheat, barley, maize and rice. Other sources include peanuts and meat products (see Pitt and Hocking, 1997). References. Pitt (1979b, 2000); Domsch et al. (1980); Samson and Frisvad (2004).

Penicillium camemberti Thom

Fig. 7.35

Penicillium candidum Roger Penicillium caseicola Bainier

Colonies on CYA 25–35 mm diam, occasionally smaller, plane or lightly radially sulcate, convex, floccose; mycelium white; conidal production usually absent to light, pale grey green or in some isolates persistently white, occasionally heavier, greyish green (25-26C3); clear exudate sometimes present; reverse pale, yellow or weakly reddish brown. Colonies on MEA 25–40 mm diam, plane, similar to those on CYA, but without exudate. Colonies on G25N 18–22 mm diam, plane or lightly radially sulcate, similar to those on MEA. At 58C, colonies commonly 3–6 mm diam. No growth at 378C. On CSN, colonies 15–20 mm diam, growing strongly, with medium reaction alkaline (purple), though often neutral near the margins; reverse alkaline. Conidiophores borne from aerial hyphae, stipes 200–400 mm long, with smooth or roughened walls, typically bearing terminal terverticillate or quaterverticillate penicilli, sometimes irregular; phialides ampulliform, 10–12(–15) mm long, with long, wide collula; conidia subspheroidal to spherical, smooth walled, 3.5–5.0 mm long, borne in short, disordered chains.

234

7

Penicillium and Related Genera

Fig. 7.35 Penicillium camemberti (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

Distinctive features. Apart from its unique habitat in cheeses and cheese factories, Penicillium camemberti is readily distinguished by its white, floccose colonies, sometimes coloured pale grey in age by tardily produced conidia. Penicilli are large and often irregular; conidia are white or grey, large and smooth walled. Strong growth occurs on CSN, with an alkaline response in medium and reverse. A violet reaction is produced with Ehrlich reagent (Lund, 1995a). Taxonomy. Raper and Thom (1949) recognised two species used for the manufacture of white cheeses, Penicillium camemberti and P. caseicola. The two species were distinguished by conidial

colour, as conidia of P. caseicola remained white in age. Later taxonomists concluded that the strains with white conidia are mutants of the grey green parent, which have been selected for properties desirable in cheese manufacture. P. camemberti is the earliest valid name for this species (Samson et al., 1977a; Pitt, 1979b). P. camemberti is correctly described as a domesticated species, derived from P. commune as the wild type (Pitt et al., 1986; Polonelli et al., 1987). Physiology. Penicillium camemberti grows strongly at refrigeration temperatures. No other information is available.

7.7

Genus Penicillium

235

Mycotoxins. Penicillium camemberti produces cyclopiazonic acid on synthetic media (Frisvad and Samson, 2004). Although most studies have reported that the toxin does not occur in cheese, not all agree. Cyclopiazonic acid is quite toxic (Purchase, 1971), is immunosuppressive in low doses (Hill et al., 1986) and has produced a mycotoxicosis in pigs (Lomax et al., 1984; Keblys et al., 2004). A role in ‘‘Turkey X’’ disease along with aflatoxin appears likely (Cole, 1986; Bradburn et al., 1994). Although no direct evidence of toxicity to humans has been reported, absence of toxicity in cheeses cannot be taken for granted, as searches for naturally non toxigenic stains have been unsuccessful (Leistner, 1990). Ecology. Penicillium camemberti and its white mutant derivatives are used in the manufacture of soft cheeses, such as Camembert, Brie and Neufchatel, and are rarely found away from the local environment surrounding the manufacture of such cheeses. P. camemberti has occasionally been isolated from other sources: meats and pecans (see Pitt and Hocking, 1997). References. Samson et al. (1977a); Pitt (1979b, 2000); Pitt et al. (1986); Samson and Frisvad (2004).

Penicillium chrysogenum Thom

Fig. 7.36

Penicillium griseoroseum Dierckx (rejected name) Penicillium notatum Westling Penicillium meleagrinum Biourge

Colonies on CYA 35–45 mm diam, occasionally less, radially sulcate, usually low and velutinous; mycelium white to yellowish; conidial production light to moderate, greyish turquoise to dull green (24-25DE3-4), in some isolates appearing more yellow green because of the presence of exudate; pale to brilliant yellow or yellow brown exudate and bright yellow soluble pigment usually produced; reverse usually brilliant yellow or yellow brown, but pale or red brown in the absence of soluble pigment. Colonies on MEA 25–40 mm diam, usually plane, low and velutinous, occasionally floccose centrally or somewhat granular; mycelium inconspicuous; conidial production moderate to heavy, greyish turquoise to dull green (24-26D3, 26-27E3-4); reverse pale, yellowish, yellow brown or reddish brown. Colonies on G25N 18–22 mm diam, usually radially sulcate and dense; reverse pale to bright yellow brown or

reddish brown. At 58C, at least microcolony formation; sometimes visible colonies up to 4 mm diam produced. At 378C, response varying from no growth to colonies up to 5 mm diam. On CSN, colonies mostly 12–18 mm diam, growth moderate to strong, medium neutral or weakly acid (yellow); reverse similar. Conidiophores borne from surface or subsurface hyphae, stipes commonly 200–300 mm long, with thin smooth walls, penicilli typically terverticillate, with 1–2 rami, either terminal and appressed or sometimes subterminal and divergent, in that case appearing biverticillate; phialides ampulliform, 7–8(–10) mm long; conidia ellipsoidal to subspheroidal, 2.5–4.0 mm long, smooth walled, borne in long, irregular columns. Distinctive features. Penicillium chrysogenum is usually a readily recognisable species: colonies grow rapidly on the standard media at 258C, and on CYA produce blue-green conidia, and yellow exudate, soluble pigment and reverse; microscopically penicilli are terverticillate, smooth walled and rather delicate by comparison with those of P. expansum or P. brevicompactum. However, some isolates lack the yellow pigmentation. Growth on CSN is moderate to strong, with a neutral or weakly acid reaction in both medium and reverse. Taxonomy. The name Penicillium griseoroseum predates P. chrysogenum (Cruickshank and Pitt, 1987; Frisvad and Filtenborg, 1989). To overcome this problem, P. chrysogenum was conserved under the provisions of the International Code of Botanical Nomenclature (Frisvad et al., 1990c; Kozakiewicz et al., 1992). Physiology. A mesophilic species, Penicillium chrysogenum has a minimum temperature for growth of 48C, an optimum at 238C and a maximum at 378C (Mislivec and Tuite, 1970b; Pitt, 1979b). Among the most xerophilic Penicillia, this species has been observed to germinate at 0.78 aw by Hocking and Pitt (1979), at 0.79 aw by Armolik and Dickson (1956) and 0.81 aw by Mislivec and Tuite (1970b). The minimum inhibitory concentration of sorbic acid effective against P. chrysogenum was reported to be 1–2 mmol/l of undissociated acid across the pH range 4–6 (Skirdal and Eklund, 1993). Ethanol (4%, w/w) inhibited the germination of P. chrysogenum conidia on PDA (Dantigny et al., 2005).

236

7

Penicillium and Related Genera

Fig. 7.36 Penicillium chrysogenum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

Penicillium chrysogenum is a major cause of tainting in foods transported in shipping containers. The taint is due to chloroanisole production from chlorinated phenols used to preserve timber in containers. P. chrysogenum was isolated from several such timber samples and shown to carry out this biochemical conversion (Hill et al., 1995). Mycotoxins. Penicillium chrysogenum produces roquefortine C, PR toxin, penicillin and secalonic acids (Frisvad and Samson, 2004). Roquefortine C

induced in vivo inflammatory responses in mouse lungs, following instillation with purified toxin. Mice subjected to high doses (12.5 nM/g of body weight of animal) of roquefortine C showed signs of trembling and lethargy until 24 h post instillation (Rand et al., 2005). However, P. chrysogenum does not appear to be a serious source of mycotoxins in foods. Ecology. Penicillium chrysogenum is a ubiquitous fungus and occupies a very wide range of habitats. As a contaminant of foods, it is probably more

7.7

Genus Penicillium

237

common even than P. aurantiogriseum. The original high penicillin producing strain of P. chrysogenum was isolated from a spoiled cantaloupe (Raper and Thom, 1949), and it has occasionally caused spoilage in stored grapes (Barkai-Golan, 1974) and carrots (Snowdon, 1991). Apart from these records, P. chrysogenum is not known as a pathogen. Few records report spoilage: that of margarine in Australia (Hocking, 1994) is an exception. This species has been reported very commonly from cereals: for example rice, wheat, barley, maize, flour (see Pitt and Hocking, 1997; Lugauskas et al., 2006) and maize-based snack foods (see Pitt and Hocking, 1997). Other major sources have been luncheon meats (Ismail and Zaky, 1999; Mohamed and Hussein, 2004), Spanish ham (Ockerman et al., 2001), dry-cured ham (Rodrı´ guez et al., 1998), cheese (Hayaloglu and Kirbag, 2007), dried fish, nuts and spices (see Pitt and Hocking, 1997). In our laboratory, we have isolated P. chrysogenum from spoiled bakery products, flavoured dairy products, cheese, margarine, lactose powder and pharmaceutical products. Several authors have looked at the possibility of using Penicillium chrysogenum as a starter culture for fermented European meat production (e.g. El-Banna et al., 1987a; Philipp and Pedersen, 1988; Krotje, 1992) and a non toxigenic strain of P. chrysogenum (Pg222) has been used as a starter culture in the ripening of dry-cured ham (Martı´ n et al., 2004, 2006). References. Samson et al. (1977b); Pitt (1979b, 2000); Domsch et al. (1980); Samson and Frisvad (2004).

Penicillium commune Thom

Fig. 7.37

Penicillium lanosum Westling Penicillium lanosogriseum Thom

Colonies on CYA 30–37 mm diam, radially sulcate, velutinous to floccose; mycelium white, usually inconspicuous; conidial production moderate, of variable colour, greyish turquoise to dull green (24-27D-F3-5); exudate usually present, clear to pale brown; reverse usually pale, occasionally yellow, brown or purple. Colonies on MEA 23–30 mm diam, plane or lightly sulcate, low and dense, surface velutinous or fasciculate; mycelium inconspicuous, white; conidial production moderate, dull green (26-27D-E3-4); reverse usually uncoloured.

Colonies on G25N 18–22 mm diam, plane, sulcate or wrinkled, low to moderately deep, dense, usually fasciculate; mycelium white to yellowish; reverse pale to orange brown. At 58C, at least microcolony formation; typically colonies of 2–4 mm diam formed. No growth at 378C. On CSN, colonies 20–26 mm diam, growth strong, medium reaction neutral to alkaline, less commonly weakly acid (pale yellow); reverse alkaline (purple). Conidiophores borne singly or in fascicles, mostly from subsurface hyphae, stipes 200–400 mm long, or of indeterminate length in fascicles, with walls finely to conspicuously roughened, typically bearing terminal terverticillate penicilli; rami 1–2 per penicillus, phialides ampulliform, 9–11 mm long; conidia spherical, less commonly subspheroidal, 3.5–4.0(–5.0) mm diam, smooth walled, borne in disordered chains. Distinctive features. Penicillium commune is similar to P. aurantiogriseum in many characteristics. However, P. commune produces grey green rather than grey blue conidia on MEA and growth of P. commune is often slower on that medium. Stipes are finely roughened to rough, only rarely smooth. P. commune is also distinguished by the ability to grow strongly on CSN and shows a positive reaction to the Ehrlich test (Lund, 1995a). Techniques for rapidly identifying this species in cheese factories were described by Lund (1996). Immunological procedures suitable for detecting Penicillium commune in P. camemberti starter cultures were developed by Polonelli et al. (1987). Differentiation of P. commune from the closely related species P. palitans and P. solitum by secondary metabolite analysis was described by Lund (1995b). Taxonomy. Regarded as a rare floccose species by Raper and Thom (1949), Penicillium commune was placed in synonymy with P. puberulum by Pitt (1979b). Pitt et al. (1986) showed that P. puberulum was a synonym of P. aurantiogriseum, so P. commune was revived as the earliest valid name for the isolates in question. P. commune is the wild type ancestor of P. camemberti (Pitt et al., 1986; Polonelli et al., 1987). Physiology. Like most other species in Penicillium subgenus Penicillium, P. commune grows rapidly at refrigeration temperatures, has an optimum near 258C and a maximum near 358C. It is probably capable of growth below 0.85 aw. P. commune failed to grow in an atmosphere of 20% CO2 and less than

238

7

Penicillium and Related Genera

Fig. 7.37 Penicillium commune (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

0.5% O2. However, it grew slowly in an atmosphere of 80% CO2 and 20% O2 (Taniwaki, 1995). Mycotoxins. Most isolates of Penicillium commune produce cyclopiazonic acid (El-Banna et al., 1987b; Polonelli et al., 1987; Frisvad and Filtenborg, 1989), a toxin described in more detail under P. camemberti. A variety of other possibly toxic compounds may also be produced, including cyclopaldic acid, cyclopolic acid, cyclopiamine, palitantin and rugulovasines (Frisvad and Filtenborg, 1989;

Lund, 1995b). P. commune produced cyclopiazonic acid, rugulovasines and viridicatin on cheese agar (Larsen et al., 2002). Ecology. The primary habitat for Penicillium commune in foods is cheese, of which it is the principal cause of spoilage (Lund et al., 1995; Kure and Skaar, 2000, 2001, 2004; Hayaloglu and Kirbag, 2007). We have isolated it frequently from spoiled soft and hard cheeses in our laboratory, as well as from yoghurt, margarine, sour cream, lactose powder and cakes

7.7

Genus Penicillium

239

containing high fat fillings. It was a major cause of ‘‘thread mould’’ spoilage of vacuum packed cheese blocks in Australia (Hocking and Faedo, 1992). The domesticated species P. camemberti probably had its origins during manufacture of cheeses over centuries. Penicillium commune has also been implicated as a cause of ‘‘phenol defect’’ in Italian hams during ripening (Spotti et al., 1988). Other sources include apples (Amiri and Bompeix, 2005), pears and associated processing machinery (Sanderson and Spotts, 1995), flour (Dragoni et al., 1980c) and European sausages ´ (Lopez-Dı´ az et al., 2001; Papagiani et al., 2007). We isolated P. commune from maize, peanuts, soybeans and cowpeas in Southeast Asia, but always at low levels (Pitt et al., 1993, 1994, 1998a). References. Pitt (1979b, 2000); Samson and Frisvad (2004).

Penicillium crustosum Thom

Fig. 7.38

Colonies on CYA 30–40 mm diam, plane or less commonly radially sulcate, typically low with a velutinous or granular texture and surface appearing powdery, sometimes with small coremia at the margins or centres; mycelium inconspicuous, white; conidial production heavy over the entire colony area, coloured predominantly dull green (26-27D-E3-4) or slightly greyer (26D2), often greyish turquoise (24D3) in marginal areas; exudate clear to pale brown or occasionally deep brown; soluble pigment sometimes present, brown; reverse pale or more commonly yellow to orange brown, often intensely coloured at the margins. Colonies on MEA 25–40 mm diam, plane, usually low and velutinous; mycelium subsurface; conidia produced very abundantly, characteristically forming masses with a dry powdery appearance, breaking off in large numbers or in crusts when jarred, coloured dull green (26–27C-D3); reverse pale or yellow brown. Colonies on G25N 20–26 mm diam, finely radially sulcate, deep but dense; yellow or brown soluble pigment occasionally produced; reverse pale, yellow brown or brown. At 58C, typically macroscopic colonies formed, 2–6 mm diam. No growth at 378C. On CSN, colonies 25–30 mm diam, showing strong growth, medium reaction acid (yellow) or occasionally alkaline (purple); colony reverse alkaline, occasionally weakly so.

Conidiophores mostly borne from subsurface hyphae, stipes commonly 200–400 mm long, with rough walls, bearing terminal penicilli, terverticillate to quaterverticillate; phialides ampulliform, 9–11 mm long; conidia spherical, less commonly subspheroidal or ellipsoidal, 3.0–4.0 mm diam, with smooth walls, borne in long parallel columns, on MEA adhering in masses. Distinctive features. Colonies of Penicillium crustosum on CYA often show blue green margins, but mature conidia en masse are definitely grey green, not bright yellow green, blue green or blue. All conidiophore elements are large, and stipe walls are definitely rough. When grown on MEA for 7–10 days, nearly all isolates of P. crustosum produce enormous numbers of conidia which readily break loose when the Petri dish is jarred. This is a remarkably consistent and useful diagnostic character. Isolates of Penicillium crustosum produced a yellow colour with the Ehrlich test (Lund, 1995a). Growth on CSN is strong, usually with an acid medium reaction and an alkaline colony reverse. Physiology. Perhaps because of poor recognition, physiological studies on Penicillium crustosum are meagre. From our data obtained during growth studies, it can be inferred that it will grow down to ca. –28C, has an optimum near 258C and a maximum about 308C. The pH limits for growth are from less than 2.2, optimally 4.5–9.0, to above 10.0 (Wheeler et al., 1991). This species is an efficient degrader of sorbic acid, producing 1,3-pentadiene, which results in off-odours in many kinds of preserved foods (Kinderlerer and Hatton, 1990). Rancidity in oils due to ketone formation is also caused by growth of fungi including Penicillium crustosum (Kinderlerer and Hatton, 1991). It is also capable of producing chloroanisoles from chlorophenols used as preservatives, leading to taints in chickens from mouldy litter (Curtis et al., 1974), in goods packed in fibreboard cartons and in paper or jute sacks (Tindale et al., 1989). Mycotoxins. Penicillium crustosum is the major producer of penitrem A, a powerful neurotoxin (Pitt, 1979c; El-Banna et al., 1987b; Frisvad et al., 2006a). Toxicity of penitrem A to animals is well documented (see Pitt and Hocking, 1997; Naude et al., 2002; Minervini et al., 2002) and it has recently been implicated in a tremor syndrome in humans (Lewis et al., 2005). All isolates of P. crustosum

240

7

Penicillium and Related Genera

Fig. 7.38 Penicillium crustosum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

produce penitrem A at high levels (Sonjak et al., 2005; Frisvad et al., 2006a), so the presence of this species in foods (or feeds) is a warning signal (Pitt, 1979c; El-Banna et al., 1987a). Penitrem A is produced only at high moisture levels, above about 0.92 aw, with an optimum around 0.995 aw (ICMSF, 1996b). This probably explains the relatively low number of reports of poisoning from a very toxic compound produced by a very common fungus.

Penicillium crustosum also produces roquefortine C and the cardiotoxin terrestric acid (Frisvad and Samson, 2004). All 120 strains of P. crustosum tested were able to produce roquefortine C, viridicatols, and terrestric acid as well as a range of other secondary metabolites (Sonjak et al., 2005). P. crustosum also produces thomitrems A and E (Rundberget et al., 2004). Ecology. Penicillium crustosum is a ubiquitous spoilage fungus. It has, for example, been isolated

7.7

Genus Penicillium

from the majority of cereal and animal feed samples examined by us over two decades. Because of confusion over species names, it was seldom mentioned in the earlier literature, but our examination of isolates published under a range of names indicates that P. crustosum has been responsible for spoilage of maize, processed meats, cheese, biscuits, cakes and fruit juices. It has been isolated as a weak pathogen from citrus fruits and melons (Snowdon, 1990) as well as apples and pears (Hee et al., 2002; Yun et al., 2006) and onions (in our laboratory). We have also isolated P. crustosum from spoiled dairy and jelly desserts, bread, rice product, fresh pistachios and from a soup can implicated in a human mycotoxicosis (Lewis et al., 2005). Other published sources include fresh cabbage, processed meats, hazelnuts, pistachios, peanuts, dried peas and amaranth grain (see Pitt and Hocking, 1997), cheese (Lund et al., 1995; Minervini et al., 2002; Hayaloglu and Kirbag, 2007) and chestnuts (Overy et al., 2003). While not commonly isolated from Southeast Asian commodities, Penicillium crustosum occasionally occurred in high numbers. Ten percent of Philippine peanut samples contained this species, with up to 40% of individual nuts infected, resulting in infection of 1% of all kernels examined. Only one of 82 maize samples from Indonesia contained P. crustosum, but more than 50% of kernels in that sample were infected. Other sources were sorghum, rice, soybeans, mung beans and pepper, at levels of 1% or less total infected particles (Pitt et al., 1993, 1994, 1998a). Additional species. Penicillium echinulatum Fassat. differs from P. crustosum by producing distinctly roughened conidia which are dark green en masse. Growth rates on the standard media are similar, but colonies of P. echinulatum usually produce copious clear to pale brown exudate, while conidia on MEA do not usually form crusts. Penicilli of the two species are similar. P. echinulatum grows strongly on CSN (22–25 mm diam). Colony reverses are usually alkaline, similar to P. crustosum, but unlike P. crustosum, an alkaline (violet) reaction is normally produced in the medium. Penicillium echinulatum has been reported to produce territrems (Frisvad and Samson, 2004). P. echinulatum is not commonly isolated from foods but has been reported from pecans, processed meats, rice, katsuobushi (see Pitt and Hocking, 1997) and margarine (in our laboratory).

241

References. Pitt (1979b; 2000); Samson and Frisvad (2004).

Penicillium digitatum (Pers.: Fr.) Sacc. Fig. 7.39 Colonies on CYA 35–55 mm diam, plane, surface texture velutinous to deeply floccose; mycelium white; conidial production moderate to heavy, greyish green to olive (1D-E3); reverse pale or brownish. Colonies on MEA of variable diameter, from 35 mm to greater than 70 mm, plane, relatively sparse, strictly velutinous; conidial production moderate, greenish olive or dull yellow green (30D4); reverse pale or brownish. Colonies on G25N 6–12 mm diam, plane, sparse, often mucoid; reverse pale or olive. At 58C, at least germination, sometimes colonies up to 3 mm diam. No growth at 378C. On CSN, colonies 4–10 mm diam, growth weak, with medium pH remaining neutral and with no change in reverse colour. Conidiophores borne from surface or aerial hyphae, stipes 70–150 mm long, with thin, smooth walls, bearing terminal penicilli, when best developed terverticillate but frequently biverticillate or irregular; phialides broadly ampulliform to cylindroidal, 10–15(–20) mm long, narrowing abruptly to large cylindroidal collula; conidia ellipsoidal to cylindroidal, 6–8(–15) mm long, smooth walled, borne in disordered chains. Distinctive features. The production of conidia coloured yellow green to olive on all substrates and the close association with rotting fruit of Citrus species distinguish Penicillium digitatum. It is also distinctive microscopically: no other species of Penicillium consistently produces such large phialides or conidia. Physiology. Penicillium digitatum can grow between 6–78C and 378C (Domsch et al., 1980). The minimum aw for growth at 258C is 0.90 (Hocking and Pitt, 1979; Plaza et al., 2003), but 0.95 aw at 308C and 0.99 at 58C. No germination was recorded at 0.87 aw or at 378C (Plaza et al., 2003). The minimum inhibitory concentration of sorbic acid preventing growth of P. digitatum was 0.02–0.025% at pH 4.7 and 0.06–0.08% at pH 5.5 (Ray-Schroeder, 1983). Mycotoxins. This species has not been reported to produce significant mycotoxins. However, about 70% of 24 isolates of Penicillium digitatum were

242

7

Penicillium and Related Genera

Fig. 7.39 Penicillium digitatum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

toxic to brine shrimp or chick embryos (Faid and Tantaoui-Elaraki, 1989). Ecology. The cause of a destructive rot of Citrus fruits (Snowdon, 1990), Penicillium digitatum is universally distributed, with a preference for warmer climates (Domsch et al., 1980). Initial control involves reducing spore build up, in the orchard by removing fallen fruit and in the factory by removing culled fruit (Snowdon, 1990). Fungicidal sprays or dips provide the main control measure, but resistance to thiabendazole, benomyl and imazalil has developed in most

countries to a greater or lesser degree (see Pitt and Hocking, 1997). Control of imazalil-resistant P. digitatum by pyrimethanil has been reported (Smilanick et al., 2006). Other control techniques include the use of essential oil vapours (Szczerbanik et al., 2007), UV light in conjunction with either a biocontrol agent or heating (D’Hallewin et al., 2005; Ben-Yehoshua et al., 2005), curing at elevated temperatures, which induces tissue healing (Plaza et al., 2004; Nunes et al., 2007) and vapour heat (Brown et al., 1991). Bacterial and yeast sprays are being

7.7

Genus Penicillium

243

developed as biocontrol agents and their commercialisation is currently been examined (Torres et al., 2007). It has been proposed that the acidification of host tissue by Penicillium digitatum enhances the pathogen’s virulence and that pH may be an important regulator of gene expression in pathogenesis (Prusky et al., 2004). Penicillium digitatum has occasionally been isolated from other food sources: hazelnuts and pistachio nuts (see Pitt and Hocking, 1997), kola nuts (Adebajo, 2000), black olives (Heperkan et al., 2006), rice, maize and meats (see Pitt and Hocking, 1997). We isolated this species at low levels from Southeast Asian peanuts, soybeans and sorghum (Pitt et al., 1993, 1994). References. Pitt (1979b, 2000); Domsch et al. (1980); Samson and Frisvad (2004).

Penicillium expansum Thom

Fig. 7.40

Colonies on CYA 30–40 mm diam, lightly radially sulcate, moderately deep to very deep, with surface typically tufted (coremial) in one or more annular bands, with adjacent areas velutinous to floccose; mycelium white; conidia produced in moderate numbers, dull green (27E3-4); exudate clear to pale orange brown; soluble pigment brownish orange near caramel (6C6); reverse pale to deep brown, often with areas of brownish orange (7C-E7-8). Colonies on MEA variable, ranging from 20 to 40 mm diam, plane, some isolates persistently velutinous, others at least partly coremial; mycelium often entirely subsurface; conidial production usually heavy, coloured as on CYA or slightly greyer (27C-D3-4); soluble pigment sometimes produced, coloured as on CYA; reverse pale or, in the presence of soluble pigment, orange brown. Colonies on G25N 17–22 mm diam, radially sulcate, dense, surface texture velutinous to granular; reddish brown soluble pigment sometimes produced; reverse pale, dull brown or reddish brown. At 58C, occasionally only microcolonies, typically colonies of 2–4 mm diam formed. No growth at 378C. Growth on CSN strong, 24–30 mm diam, medium acid (yellow); reverse usually acid plus brown soluble pigment, occasionally alkaline (purple). Conidiophores borne from surface or subsurface hyphae, singly, in fascicles, or in definite coremia, sometimes visible with the unaided eye, stipes 200–500 mm long, with smooth walls, bearing terminal penicilli,

typically terverticillate, less commonly biverticillate; rami borne singly, phialides closely packed, ampulliform to almost cylindroidal, 8–11 mm long, with short collula; conidia ellipsoidal, 3.0–3.5 mm long, smooth walled, borne in long, densely packed, irregular chains. Distinctive features. The most important features distinguishing Penicillium expansum are dull green conidia, often borne in coremia, smooth walled stipes on both CYA and MEA and, frequently, the presence of orange brown to brown exudate, soluble pigment and reverse colours. Isolates of P. expansum inoculated into apples or pears typically produce destructive rots. Growth on CSN is strong; medium and reverse colours are typically yellow. P. expansum produces a violet colour with Ehrlich reagent (Lund, 1995a). Physiology. Like most other species in this subgenus, Penicillium expansum is a psychrophile: minimum temperatures for growth have been reported as –68C (Brooks and Hansford, 1923), –38C (Panasenko, 1967) and at most –28C (Mislivec and Tuite, 1970b). Growth is quite strong at 08C (Kuehn and Gunderson, 1963). The optimum temperature for this species is near 258C and the maximum near 358C (Panasenko, 1967). The effect of temperature on the germination time and growth rate of Penicillium expansum has been modelled by Baert et al. (2007a). They also provided excellent data for the optimum temperature for growth being 258C. The minimum aw for germination is 0.82–0.83 (Mislivec and Tuite, 1970b; Hocking and Pitt, 1979). Sodium chloride was more inhibitory than glucose, glycerol or sorbitol at aw values between 0.98 and 0.89, with germination and growth at 158C only slightly slower than at 258C (Lahali et al., 2005). At 58C, growth ceased at 0.91 aw (NaCl and glucose) or 0.89 aw (glycerol, sorbitol) but the incubation period was only 25 days (Lahali et al., 2005). Penicillium expansum has a very low requirement for oxygen. Golding (1940a, 1945) showed that growth was virtually unaffected by levels of oxygen as low as 2.1%. When reduction in rates of growth did occur, it was at higher temperatures. Growth of P. expansum and some other fungi was stimulated by carbon dioxide concentrations up to 15% in air, but growth rates declined at higher CO2 levels. Spores of P. expansum were inactivated by holding in an atmosphere of 13% CO2 for 21 days, assisting the fumigation of fruit boxes (Cossentine et al., 2004).

244

7

Penicillium and Related Genera

Fig. 7.40 Penicillium expansum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

Mycotoxins. Penicillium expansum is an important producer of patulin and citrinin (Harwig et al., 1973; Ciegler et al., 1977). Patulin is reported to affect renal cells (Heussner et al., 2006) and decrease sperm count in rats (Selmanog˘lu, 2006). Patulin damaged the DNA of mammalian cells, but the effect could be negated by ascorbic acid (Alves et al., 2000). Patulin also caused oxidative stress response in mammalian cell lines, which can lead to cell changes during transformation and differentiation (Liu et al., 2007). However, patulin is still classified as a Group 3

compound by the International Agency for Research on Cancer (IARC) on the basis that cytotoxicity in animals does not constitute adequate evidence of human toxicity (IARC, 1993). Despite a lack of definite data on human toxicity, a number of countries have set an upper limit of 50 mg/l for patulin in apple juice and other apple products (FAO, 2003), with a limit of 10 mg/l for apple products destined for consumption by children (EU, 2004). Penicillium expansum produces patulin as it rots apples and pears. Poor quality control, i.e. the use of

7.7

Genus Penicillium

rotting fruit in juice or cider manufacture can result in high concentrations of patulin in juice (Brackett and Marth, 1979; Watkins et al., 1990). Levels of patulin ranging from 538 to 1822 mg/ml were detected in apple cider stored at 258C and up to 396 mg/ml in cider stored at 48C (McCallum et al., 2002). Aggressively pathogenic strains of P. expansum produced more patulin than less aggressive strains, and patulin synthesis seemed correlated with an ability to increase the acidity of cider, a characteristic of the virulent strains (McCallum et al., 2002). Patulin is important as an indicator of the use of poor quality raw materials in juice manufacture. Patulin can be produced over the range 0–258C at least, the optimum being at 258C (Paster et al., 1995), but not at 318C (Northolt et al., 1978). The minimum aw for patulin production by Penicillium expansum is 0.95 at 258C (Patterson and Damoglou, 1986). Patulin is produced over the narrow pH range of 3.2–3.8 in apple juice (Damoglou and Campbell, 1986). A modified atmosphere of 3% CO2 and 2% O2 completely inhibited patulin production at 258C, but production occurred in atmospheres of 2% CO2 and 10 or 20% O2 (Paster et al., 1995). Patulin in quite stable in apple juice during storage (Damoglou and Campbell, 1986), and pasteurisation at 908C for 10 seconds caused less than 20% reduction (Wheeler et al., 1987). No patulin was produced in apples stored under controlled atmospheres of either 2.5% O2/3.9% CO2 or 1.5% O2/2.5% CO2 at 18C. Patulin was only detected in apples after further storage at 208C (Morales et al., 2006, 2007a). It has been shown that Penicillium expansum invades apples through the stems during storage (Rosenberger et al., 2006). Ecology. One of the oldest described Penicillium species, P. expansum has been established as the principal cause of spoilage of pome fruits throughout this century (Raper and Thom, 1949). Isolates of P. expansum come predominantly from rotting apples and pears (Snowdon, 1990). Control relies on sprays to prevent Colletotrichum infections which permit entry by P. expansum, prevention of damage to fruit, cool storage and rapid processing (Snowdon, 1990). A wide variety of other techniques have been advocated, including use of deoxyglucose as a control agent (Janisiewicz, 1994; El-Ghaouth et al., 1995), chlorine dioxide in wash waters (Roberts and

245

Reymond, 1994; Okull et al., 2006) or wash water filtration (Spotts and Cervantes, 1993) and calcium chloride infiltration under increased temperature and pressure (Conway et al., 1988; Sams et al., 1993). Biological control using bacteria or yeasts has also been studied, and two commercial products, one based on Candida oleophila and the other on Cryptococcus albidus have been registered in the United States and South Africa (Janisiewicz and Korsten, 2002; Fravel, 2005). However, the efficacy of these products still depends on supplementation with low doses of synthetic fungicides to achieve large scale control (Droby et al., 2003). Other compounds including salicylic acid and cytokinin have been examined in conjunction with biocontrol yeasts to enhance the control of P. expansum in pears (Yu et al., 2007; Zheng et al., 2007b). Penicillium expansum has been isolated from a wide range of other fruits including tomatoes, strawberries, avocados, mangoes, grapes (see Pitt and Hocking, 1997; Snowdon, 1990) indicating that it is a broad spectrum pathogen on fruits. Isolation from fresh vegetables has been uncommon: onions, carrots and cabbages (Lugauskas et al., 2005) have been reported. Isolations from stored foods have been less frequent: in particular, Penicillium expansum appears to be much less common on cereals than some other species in this subgenus. Isolations have been reported from maize, wheat, rice and barley (Aziz et al., 2006) and a variety of retail cereal products (Aran and Eke, 1987). It is more widespread in other foods, especially meat and meat products (see Pitt and Hocking, 1997 and Cantoni et al., 2007). Other records include pecans and pistachios (see Pitt and Hocking, 1997), peanuts, (Aziz et al., 2006), olives (Kivanc¸ and Akguel, 1990), dried beans, beans (see Pitt and Hocking, 1997), cheese (Hayaloglu and Kirbag, 2007) and margarine (Hocking, 1994), health foods, rapeseed, dried fish and frozen fruit pastries (see Pitt and Hocking, 1997). We have isolated P. expansum from spoiled cheese, fruit yoghurt, jellied fruit desserts, apple sauce and apple juice. Only low levels were found in Southeast Asian commodities (Pitt et al., 1993, 1994). References. Pitt (1979b, 2000); Domsch et al. (1980); Samson and Frisvad (2004).

246

Penicillium griseofulvum Dierckx

7

Fig. 7.41

Penicillium patulum Bainier Penicillium urticae Bainier

Colonies on CYA 20–25 mm diam, occasionally 30 mm, finely radially sulcate, moderately deep, dense, surface texture granular; mycelium white; conidial production moderate to heavy, at the margins greyish green (26-27C3), centrally greenish grey (26-27C2); exudate usually present, clear to pale yellow; soluble pigment sometimes produced, reddish brown; reverse pale, dull yellow or brown. Colonies on MEA 15–25 mm diam, plane or rarely radially sulcate, moderately deep, of granular texture; mycelium usually inconspicuous, white; conidial production moderate to heavy, coloured as on CYA; reverse pale to brown. Colonies on G25N 16–22 mm diam, plane, low and velutinous at the margins, often floccose centrally; reverse pale. At 58C, colonies up to 4 mm diam usually formed. No growth at 378C. On CSN, colonies 18–24 mm diam, growth moderate; medium and reverse reaction neutral. Conidiophores borne in fascicles, with stipes of indeterminate length, often sinuous, smooth walled, brownish, terminating in distinctive penicilli, sometimes terverticillate, more commonly quaterverticillate and not infrequently with five or even more branch points between stipe and phialide; phialides closely packed, exceptionally short, 4.5–6.0 mm, abruptly tapering to short collula; conidia ellipsoidal, 3.0–3.5 mm long, smooth walled, borne in closely packed, disordered chains. Distinctive features. Penicillium griseofulvum is unique in two respects: it produces very short phialides and it bears them on highly branched conidiophores. Other features include the production of grey colonies on CYA and MEA, with only weak greenish overtones, and a neutral reaction on CSN. Physiology. Growth temperatures for Penicillium griseofulvum range from 4 to 358C, with an optimum near 238C (Mislivec and Tuite, 1970b). The minimum aw for germination is 0.81 at 238C and 0.83 at 16 or 308C (Mislivec and Tuite, 1970b). Mycotoxins. Penicillium griseofulvum, sometimes reported under its synonyms P. patulum and P. urticae, produces four mycotoxins, patulin, cyclopiazonic acid, roquefortine C and griseofulvin

Penicillium and Related Genera

(Samson and Frisvad, 2004). Patulin and cyclopiazonic acid are moderately toxic compounds, with oral LD50 values in male rats and mice of 35 and 36 mg/kg, respectively. Griseofulvin, with an intravenous LD50 in rats of only 500 mg/kg, is sufficiently nontoxic to permit use as an antifungal antibiotic. Griseofulvin has been of great value in the treatment of cutaneous infections by dermatophytic fungi in both man and animals. However, as well as inhibiting invasive fungi, it also shows mammalian toxicity, acting as a mitotic blocker in the host’s dividing cells, such as those of bone marrow, the intestinal lining, and tumours (see Vanden Bossche et al., 2003 and Woodward, 2005 for reviews). When cyclopiazonic acid produced from Penicillium griseofulvum was fed to lactating ewes, most was excreted in milk within 10 days, the maximum concentration in milk (570 mg/kg) being ca. 10% of the ingested dose (5 mg/kg live weight). Appreciable amounts of this compound fed to chickens were also detected in eggs (Dorner et al., 1994). Although one outbreak of cattle poisoning in Japan was attributed to Penicillium griseofulvum (synonym P. maltum M. Hori and T. Yamamoto) (Hori et al., 1954), recent literature provides no other example of animal toxicity due to this species. Cyclopiazonic acid was reported to be non-mutagenic to human liver cells and reduced the mutagenicity of aflatoxin B1 (Sabater-Vilar et al., 2003b). It is unlikely that P. griseofulvum is a significant contributor to human illness. Patulin was produced by Penicillium griseofulvum down to 0.88 aw (Lotzsch and Trappe, 1978). ¨ Production occurred over the range 4–318C at 0.99 aw, and 8–318C at 0.95 aw (Northolt et al., 1978). Potato dextrose broth supplemented with manganese optimised patulin synthesis (Dombrink-Kurtzman and Blackurn, 2005). Ecology. Although Penicillium griseofulvum can produce lesions when inoculated into apples and pears, it is not a common cause of spoilage of these fruits (Sanderson and Spotts, 1995). P. griseofulvum occurs quite commonly on cereals and nuts. Records from cereals include barley, maize, rice and wheat (Aziz et al., 2006), flour and bakery products (see Pitt and Hocking, 1997) and from nuts: peanuts (Aziz et al., 2006), pecans and pistachios (see Pitt and Hocking, 1997). Other sources include dried peas and beans,

7.7

Genus Penicillium

247

Fig. 7.41 Penicillium griseofulvum (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) conidia, bar = 5 mm

meats, rapeseed, health foods and frozen fruit pastries (see Pitt and Hocking, 1997). P. griseofulvum occurs only rarely in tropical foods: we isolated it only from Thai and Philippine peanuts at low levels (Pitt et al., 1993, and our unpublished data). References. Samson et al. (1976); Pitt (1979b, 2000); Domsch et al., 1980); Samson and Frisvad (2004).

Penicillium hirsutum Dierckx

Fig. 7.42

Penicillium corymbiferum Westling Penicillium verrucosum var. corymbiferum (Westling) Samson et al.

Colonies on CYA 30–40 mm diam, radially sulcate or less commonly plane, surface texture typically granular to coremial; mycelium white at the margins, white to bright yellow elsewhere; conidial

248

production moderate, greyish green to dull green near Spanish green or jade green (26-27D-E4-5); exudate usually violet brown near maroon (11EF8) but occasionally lighter brown; soluble pigment typically produced, deep yellow to orange brown, occasionally reddish brown; reverse in similar colours. Colonies on MEA 25–35 mm diam, plane or rarely sulcate, surface texture usually granular to coremiform, less commonly velutinous to fasciculate; mycelium white to deep

7

Penicillium and Related Genera

yellow; conidial production moderate, in colours similar to those on CYA; exudate produced by some isolates, deep red; reverse pale to deep yellow, green or orange brown. Colonies on G25N 18–22 mm diam, plane or sulcate, low to deeply floccose; mycelium white to yellow; yellow soluble pigment often produced; reverse bright yellow green or orange to brown. At 58C, colonies of 2–5 mm diam usually produced. No growth at 378C.

Fig. 7.42 Penicillium hirsutum (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) conidia, bar = 5 mm

7.7

Genus Penicillium

On CSN, colonies 24–30 mm diam, growth strong, medium reaction acid (yellow); reverse acid, sometimes with brown pigment also. Conidiophores commonly borne from surface hyphae, with stipes 200–500 mm long, but also borne in fascicles and then of indeterminate length, typically with conspicuously roughened walls, bearing large terminal terverticillate to quaterverticillate penicilli; rami 1–2 per penicillus, typically with walls roughened; metulae usually rough walled; phialides ampulliform, smooth walled, 9–12 mm long, abruptly tapering to generally short collula; conidia spherical, 3.0–3.5 mm diam, less commonly ellipsoidal, 3.0–3.5 mm long, with walls smooth to very finely roughened, borne in disordered chains or irregular columns. Distinctive features. On CYA, colonies of Penicillium hirsutum are usually deep and fasciculate to coremiform, exudate is reddish or violet brown, soluble pigment is deep yellow to orange brown and conidia are usually pure green, without blue or yellow tones. Conidiophores are frequently borne in fascicles, stipes are usually rough walled and penicilli are large. P. hirsutum grows strongly on CSN, and normally produces an acid reaction in both medium and colony reverse. Taxonomy. Raper and Thom (1949: 544) called this taxon Penicillium corymbiferum. Samson et al. (1976) reduced this species to varietal status under P. verrucosum, but this change in rank was not accepted by Pitt (1979b), who took up the earlier name P. hirsutum. This has now been accepted (Frisvad and Filtenborg, 1989; Pitt and Samson, 1993). Physiology. Penicillium hirsutum is capable of growth at particularly low temperatures, with a minimum of –58C and relatively rapid growth at –18C (Lowry and Gill, 1982). Overy et al. (2005a) reported growth at 08C with an optimum at 208C. It appears to be moderately xerophilic (Gill and Lowry, 1982). Mycotoxins. This species produces roquefortine C (Frisvad and Filtenborg, 1989; Frisvad and Samson, 2004). The production of cyclopiazonic acid was reported by El-Banna et al. (1987b), but this was not confirmed by later studies (Frisvad, 1989; Frisvad and Filtenborg, 1989; Frisvad and Samson, 2004). Ecology. Although an infrequently encountered species, Penicillium hirsutum is important as a cause of black spot in refrigerated meat (Gill et al., 1981). It has also caused spoilage of fresh asparagus (Saito et al.,

249

2003), onion and garlic bulbs (Overy et al., 2005b) and refrigerated pear puree (Pitt, 1979b: 351). It has been reported from wheat, flour, rice and peanuts (see Pitt and Hocking, 1997). We isolated it infrequently from Indonesian soybeans (Pitt et al., 1998a). Additional species. Penicillium allii Vincent and Pitt [synonym P. hirsutum var. allii (Vincent and Pitt) Frisvad] is similar to P. hirsutum in many respects. However, colonies are larger at 258C (35–45 mm diam on both CYA and MEA) and growth does not occur at 58C. Colonies are velutinous or minutely fasciculate. Stipes are very rough. Growth on CSN is weak, 15–20 mm diam, with an acid reaction in both medium and colony reverse. P. allii produces roquefortine C (Overy et al., 2005b). This species causes a destructive rot in garlic (Vincent and Pitt, 1989; Snowdon, 1991, as P. corymbiferum). We isolated it from 2% of all kemiri nuts examined from Indonesia (Pitt et al., 1998a), and at low levels from Philippine maize (our unpublished data). Additional species. Penicillium hordei Stolk was distinguished from P. hirsutum by the production of yellow coremia on MEA, by stipes with walls less conspicuously roughened, and by smaller, roughwalled conidia (Stolk, 1969). Pitt (1979b) considered these taxa to be a single species, while P. hordei was treated as P. hirsutum var. hordei (Stolk) Frisvad by Frisvad and Filtenborg (1989). Ecological evidence (Stolk, 1969; Frisvad and Filtenborg, 1989) and differences in isoenzyme patterns (Cruickshank and Pitt, 1987) indicated that P. hirsutum and P. hordei are separate species, and this is now accepted (Pitt and Samson, 1993; Frisvad and Samson, 2004). P. hordei isolates always produce the minor toxins roquefortine C and terrestric acid (Frisvad and Filtenborg, 1989; Overy et al., 2005b). On CSN, growth is similar to that of P. hirsutum, colonies 20–24 mm diam, growth strong, medium reaction acid, reverse acid with deep brown pigments also. This species is of quite common occurrence in barley in Europe (Stolk, 1969) and also occurs on tomatoes and in spices (Frisvad and Filtenborg, 1989). P. hordei also causes blue rot in tulip bulbs (Overy et al., 2005b). References. Penicillium hirsutum: Pitt (1979b, 2000); Samson and Frisvad (2004). P. allii: Vincent and Pitt (1989); Samson and Frisvad (2004). P. hordei: Stolk (1969); Samson and Frisvad (2004).

250

Penicillium italicum Wehmer

7

Fig. 7.43

Colonies on CYA 30–40 mm diam, plane or radially sulcate, usually low and dense, velutinous to granular, some isolates with minute coremia at the margins; mycelium white; conidia abundant, greyish green (25-26C2-3); clear exudate and brown soluble pigment produced by some isolates; reverse usually brownish orange to greyish brown (7C7-F3). Colonies on MEA 35–55 mm diam, plane and sparse, usually strictly velutinous, sometimes with minute coremia at the margins; conidial formation moderate to heavy, coloured as on CYA; reverse typically chocolate brown (6E-F4). Colonies on G25N 12– 17 mm diam, plane or sulcate; brown soluble

Penicillium and Related Genera

pigment sometimes produced; reverse yellow brown to deep brown. At 58C, microcolonies to colonies of 4 mm diam produced. No growth at 378C. On CSN, colonies 10–20 mm diam, growth weak, medium and colony reverse reaction neutral. Sclerotia produced by some isolates, up to 300 mm diam, brown and soft. Teleomorph unknown. Conidiophores borne from surface or subsurface hyphae, stipes commonly 200–400 mm long, with thin, smooth walls, bearing large regular to irregular terminal terverticillate penicilli; rami 1–2 per penicillus, metulae often apically inflated; phialides 10–14 mm long, roughly cylindroidal in shape, then tapering abruptly to long cylindroidal collula;

Fig. 7.43 Penicillium italicum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

7.7

Genus Penicillium

conidia borne as cylinders, enlarging and rounding with maturation, ellipsoidal to short cylindroidal, 3.0–5.0 mm long, with smooth walls, borne in long, disordered chains. Distinctive features. Penicillium italicum is readily recognised in nature as the cause of a destructive bluish grey rot on lemons or other Citrus fruit. In culture it forms relatively broad, grey green colonies with deep brown reverse colours, with conidia borne as cylinders, enlarging and rounding during maturation. It grows poorly on CSN, and the medium shows little or no change. Physiology. According to Panasenko (1967), Penicillium italicum grows between –38C and 32–348C, with an optimum at 22–248C; a minimum of 08C was reported by Wyatt et al. (1995). The minimum aw for germination at 10 and 258C is 0.87(Panasenko, 1967; Plaza et al., 2003). The pH range for growth is 1.6–9.8 (Panasenko, 1967). Mycotoxins. Although Penicillium italicum has been shown to exhibit toxicity in some biological assays (Faid and Tantaoui-Elaraki, 1989) and an isolate was somewhat toxic to ducklings (Kriek and Wehner, 1981), mammalian toxicity has not been shown. Ecology. The primary habitat for Penicillium italicum is fruit of Citrus species, on which it produces a destructive rot of considerable economic importance (Snowdon, 1990). Control measures are essentially similar to those used against P. digitatum. P. italicum has also shown resistance to commonly used fungicides (Wild, 1983; Diaz-Borras et al., 1987; Dave´ et al., 1989). It has been reported only rarely from other foods: from avocados, tomatoes, sapodillas and rice (see Pitt and Hocking, 1997), meat (Papagianni et al., 2007), sausages, salami and cheese (Guillet et al., 2003) and fruit juices (Wyatt et al., 1995). Additional species. Penicillium ulaiense H.M. Hsieh et al. is closely related genetically to P. italicum (Holmes et al., 1994; Frisvad and Samson, 2004). It differs by much slower growth on CYA (15–20 mm in 7 days at 258C) and MEA (15–17 mm). Colony appearance is very similar, though reverse colours on CYA and MEA are yellowish orange, and coremium production more marked (Holmes et al., 1994; Frisvad and Samson, 2004). Colonies on CSN are 4–8 mm diam, growth weak and sparse, medium and reverse neutral. Like

251

P. italicum, P. ulaiense is pathogenic on Citrus fruits, especially oranges and lemons. It has been isolated from most citrus growing areas around the world, including North and South America, southern Europe, Denmark, South Africa, Taiwan, Israel, Turkey and Australasia (Holmes et al., 1994; Carrillo, 1995; Frisvad and Samson, 2004). Most isolates are resistant to imazalil, the chemical of choice for control of P. italicum. P. ulaiense is a weaker pathogen than P. italicum (Holmes et al., 1994): they postulated that P. ulaiense is a rare species, but has developed imazalil resistance and become significant once P. italicum was controlled. Penicillium ulaiense grows quite strongly at 58C and reaches its optimum near 258C; growth is weak at 308C and absent at 338C (Holmes et al., 1994). It is not known to produce mycotoxins and has not been found away from Citrus fruits or citrus packing houses. References. Penicillium italicum: Samson et al. (1976); Pitt (1979b, 2000); Domsch et al. (1980); Samson and Frisvad (2004). P. ulaiense: Holmes et al. (1994); Samson and Frisvad (2004).

Penicillium nalgiovense Laxa

Fig. 7.44

Colonies on CYA usually 28–35 mm diam, radially sulcate, dense and velutinous to quite floccose, conidial production light to heavy, white to dull green (25-26B-E3-5); clear exudate produced by most isolates; reverse pale to bolden or blonde (4-5B-E3-6). Colonies on MEA variable, 10–30 mm diam, plane, dense to somewhat floccose, usually heavily sporing, white to dull green (26-27D-E3-5); exudate and soluble pigment absent; reverse usually strongly coloured, in deep orange brown to brown shades (6C-E6-8). Colonies on G25N 14–22 mm diam, plane, dense to floccose, conidial production moderate to heavy, white to dull green, as on CYA; exudate and soluble pigment absent; reverse pale, yellow or golden brown. At 58C, germination. No growth at 378C. Colonies on CSN usually 10–18 mm diam, of neutral to weakly acid reaction in both medium and colony reverse. Conidiophores borne from surface hyphae, stipes commonly 150–400 mm long, or of indeterminate length, often branching irregularly from basal hyphae, smooth walled, bearing irregular penicilli; penicilli very variable within a single preparation, in

252

7

Penicillium and Related Genera

Fig. 7.44 Penicillium nalgiovense (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

the most characteristic form a compact terminal terverticillate structure with a divergent subterminal ramus, occasionally quaterverticillate or more complex, less commonly biverticillate, sometimes with intercalary irregular metulae or rami; phialides ampulliform to acerose or nearly cylindrical, 7–10 mm long, with short, broad collula; conidia spherical to subspheroidal, 2.8–3.5 mm diam, smooth walled, borne in short columns. Distinctive features. Closely related to Penicillium chrysogenum, this species is distinguished by moderate growth, white to pale green sporulation, usually deep orange reverse colours on MEA, and complex, often irregularly branched penicilli. Growth on CSN is weak.

Taxonomy. For a long time, the name Penicillium nalgiovense has been applied to a mould used in the manufacture of fermented meats in Europe (Incze and Mihalyi, 1976; Dragoni and Marino, 1979). Based on the type isolate, from cheese, this species was synonymised with P. jensenii by Pitt (1979b). However, comparative studies of isolates from meats with the type of P. nalgiovense have shown that all belong to a single species (R.H. Cruickshank and J.I. Pitt, unpublished). It appears likely that P. nalgiovense is a domesticated form of the common spoilage species P. chrysogenum, with a major habitat in fermented meat products in Europe. Isolates of Penicillium nalgiovense show substantial variation in morphology and the species can be

7.7

Genus Penicillium

divided into six biotypes. These vary in rates of growth on MEA and colour and degree of sporulation on CYA and MEA (Fink-Gremmels et al., 1988). Physiology. As with cheese starter cultures, much is known about the enzymatic and functional properties of strains of this species for use as starter cultures. Little has been published on the gross physiological properties of interest here. A close similarity to Penicillium chrysogenum is to be expected. Mycotoxins. The possibility of mycotoxin production by Penicillium nalgiovense has been widely investigated. Most isolates show very low toxicity, and selection of non toxigenic strains has been successful (Leistner and Pitt, 1977; Fink-Gremmels et al., 1988; Hwang et al., 1993a; Andersen, 1995). Some isolates produce penicillin (Andersen and Frisvad, 1994; Papagianni et al., 2007) and nalgiovensin. However, the toxicity of the latter is unknown (Frisvad and Samson, 2004). Diaportins have been produced by isolates of P. nalgiovense on cheese agar (Larsen et al., 2002). Ecology. The main ecological niche occupied by Penicillium nalgiovense is as a starter culture for fermented meat production in Europe. It is a reasonable postulate that this species evolved from P. chrysogenum during the millennia that such products have been made. Consult the review by Sunesen and Stahnke (2003) for the supply, use and safety of such starter cultures. Other sources are rare: they include nuts (Sahin and Kalyoncuoglu, 1994) and cheese (Lund et al., 1995). References. Fink-Gremmels et al. (1988); Samson et al. (1995); Samson and Frisvad (2004).

Penicillium olsonii Bainier & Sartory Fig. 7.45 Colonies on CYA 30–40 mm diam, plane or lightly sulcate, deep but velutinous; mycelium inconspicuous, white to pale brown; conidial production moderate to heavy over the whole colony area, greyish green to dull green (26-27C-E3); limited amounts of clear exudate sometimes produced; reverse pale yellow or yellow brown. Colonies on MEA 25–35 mm diam, plane and velutinous; mycelium inconspicuous except sometimes centrally, white to buff; conidial production moderate to heavy, greenish grey to greyish green (26-28C2-3); reverse pale or dull yellow brown. Colonies on G25N 22–28 mm diam, similar in morphology and colouration to colonies on CYA;

253

reverse pale to yellow. At 58C, germination of some conidia to formation of microcolonies. No growth at 378C. On CSN, colonies 6–14 mm diam, growth weak and sparse, no reaction in medium or colony reverse. Conidiophores borne from subsurface hyphae, very large, stipes typically 500–2000 mm long, and 4.0–6.0 mm wide, but occasionally even larger, smooth walled, terminating in characteristic closely appressed multiramulate penicilli, usually terverticillate but sometimes quaterverticillate; rami commonly 2–3 but up to 5–6 per stipe; phialides ampulliform, 9–10 mm long, with short collula; conidia ellipsoidal, 3.0–4.0 mm long, with walls smooth or finely roughened, borne in disordered chains. Distinctive features. Exceptionally large multiramulate terverticillate penicilli borne on long, wide stipes set Penicillium olsonii apart from other Penicillium species. Average penicilli probably contain some 200 phialides and numbers in excess of 300 are not impossible. Taxonomy. Penicillium olsonii appears to be closely related to P. brevicompactum (Stolk and Samson, 1985; Cruickshank and Pitt, 1987) and the new species P. astrolabium isolated from grapes in Portugal (Serra and Peterson, 2007). Physiology. No physiological studies are known. Mycotoxins. No mycotoxins are produced (Frisvad and Filtenborg, 1989) but penicillin production has recently been reported (Papagianni et al., 2007). Ecology. This was a very poorly recognised species until recently, so reports of its occurrence in foods are very limited. It is not a rare species, however. It comprised 8–15% of isolates from a large number of mould ripened sausages (Andersen, 1995; Papagianni et al., 2007). We have seen it occasionally from spoiled margarine in our laboratory. We isolated it from 20% of soybean samples from the Philippines; it was present on up to 34% of individual kernels in a sample, and on 3% of all seeds examined (our unpublished data). It occurred at low levels in maize, peanuts, cashews, sorghum, mung beans and copra from Thailand, from maize and paddy rice in the Philippines (our unpublished data) and from peanuts and mung beans in Indonesia (Pitt et al., 1993, 1994, 1998a). References. Pitt (1979b, 2000); Samson and Frisvad (2004).

254

7

Penicillium and Related Genera

Fig. 7.45 Penicillium olsonii (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

Penicillium roqueforti Thom

Fig. 7.46

Colonies on CYA and MEA growing rapidly, 40– 70 mm diam, plane or lightly radially sulcate, low, strictly velutinous; mycelium inconspicuous, white; conidial production moderate to heavy, at the margins greyish turquoise (24C3-4), predominantly Dull Green (25-26E4) and sometimes centrally Olive Brown (4D3-4); reverse pale, brown, or green to deep blue green, almost black. Colonies on G25N usually 20–22 mm diam, but sometimes with

spreading submerged margins and then up to 28 mm diam, plane or lightly radially sulcate, colours similar to those on CYA. At 58C, colonies usually 2–5 mm diam. No growth at 378C. On CSN, colonies 25–40 mm diam, growth strong (though not deep), medium reaction variable, from acid (yellow) to alkaline (violet); reverse usually alkaline. Conidiophores borne from subsurface hyphae, stipes 100–200 mm long, with walls characteristically very rough, bearing large terminal penicilli, typically

7.7

Genus Penicillium

255

Fig. 7.46 Penicillium roqueforti (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) conidia, bar = 5 mm

terverticillate, occasionally quaterverticillate or rarely biverticillate, with elements appressed; rami rough walled; metulae occasionally rough walled; phialides ampulliform, commonly 8–10 mm long, with short collula; conidia spherical, 3.5–4.0(–6) mm diam, with thin, perfectly smooth walls, dark green, borne in long, closely packed, irregular to disordered columns.

Distinctive features. Penicillium roqueforti grows very rapidly and produces low and velutinous dark green colonies; stipes have very rough walls and conidia are large, spherical and smooth walled. Growth on CSN is rapid and colony reverse is usually alkaline. Taxonomy. The species described as Penicillium roqueforti by Pitt (1979b, 2000) and Pitt and

256

Hocking (1997) comprises more than one taxon (Frisvad and Samson, 2004). Two varieties were recognised by Frisvad and Filtenborg (1989): P. roqueforti var. roqueforti, used in cheese manufacture, and P. roqueforti var. carneum, a spoilage fungus. These varieties differed in mycotoxin production and to some extent in ecology. On the basis of substantial differences in ITS DNA, Boysen et al. (1996) separated P. roqueforti into three species: P. roqueforti, P. carneum and P. paneum. Subsequent metabolic analyses have confirmed the existence of three separate species (Karlshøj and Larsen, 2005). These three species are difficult to distinguish morphologically. However, P. roqueforti isolates produce a deep green colour in the reverse on CYA, whereas the reverses of the other two species are yellow brown. Penicillium roqueforti and P. carneum produce a violet colour with the Ehrlich reaction, whereas P. paneum does not (Frisvad and Samson, 2004). P. carneum colonies on CYA produce a strong earthy smell, whereas those of P. paneum do not (Frisvad and Samson, 2004) Physiology. Penicillium roqueforti appears to have the lowest oxygen requirement for growth of any Penicillium species. It grows normally down to 75 mmHg pressure, i.e. only 10% of standard atmospheric pressure (Golding, 1940a). Growth of P. roqueforti is little affected by oxygen concentrations down to 4.2% and is stimulated by carbon dioxide concentrations up to 15% in air (Golding, 1940b, 1945). In an atmosphere with less than 0.5% O2, P. roqueforti grew in the presence of 20% but not 40% CO2 at 258C, and growth and sporulation also occurred in an atmosphere containing 20% O2 and 80% CO2 (Taniwaki, 1995). P. roqueforti grew on rye bread in a modified atmosphere comprising 0.03% O2 and almost 100% CO2 (Suhr and Nielsen, 2005). These properties are undoubtedly a major reason for the dominant growth of this species in ripening cheeses. Like other species in this subgenus, Penicillium roqueforti is a psychrophile. It grows vigorously at refrigeration temperatures, but not above 358C (Moreau, 1980). At aw 0.89 and 0.92, conidia of P. roqueforti germinated at 25 and 308C but not at 378C or at lower aw values (Gock et al., 2003). It is tolerant of alkali, the pH range for growth being 3–10 (Moreau, 1980). Furthermore it is highly tolerant of weak acid preservatives, being able to grow in the

7

Penicillium and Related Genera

presence of 0.5% acetic acid, a property used as the basis for a selective medium (Engel and Teuber, 1978). Growth was stimulated in the presence of 0.3% propionic acid at 0.97 aw (Suhr and Nielsen, 2004). Tolerance to sorbic acid has also been observed (Liewen and Marth, 1984; Bullerman, 1984 and our unpublished data), accompanied by degradation of the preservative (Finol et al., 1982) and development of a ‘‘kerosene’’ taint due to 1,3-pentadiene formation (Liewen and Marth, 1985; Daley et al., 1986). P. roqueforti also causes taints in bread due to production of a mixture of 2-methylisoborneol and various 8carbon alcohols and ketones (Harris et al., 1986). Mycotoxins. The cheese mould Penicillium roqueforti produces PR toxin in pure culture (Frisvad et al., 2006a; Nielsen et al., 2006). It is sometimes produced in very low levels in cheese (Scott, 1981; Teuber and Engel, 1983; Schoch et al., 1984), where the toxin is degraded (Chang et al., 1993; Siemens and Zawistowski, 1993). It was not detected in Tulum cheese, a traditional Turkish cheese (Erdogan and Sert, 2004) or blue cheese in Europe (Finoli et al., 2001). P. roqueforti also produces roquefortine C and mycophenolic acid (Frisvad and Samson, 2004; Nielsen et al., 2006), but these compounds have very low toxicity (Scott, 1981). Ecology. Although best known for its role in the manufacture of Roquefort and related cheese types, Penicillium roqueforti is in fact a widely distributed spoilage fungus (though some reports may actually be P. carneum or P. paneum). Its ability to grow rapidly at refrigeration temperatures makes it a common cause of spoilage in cool stored foods, both commercial and domestic. Like P. commune, it is a common cause of cheese spoilage (see Pitt and Hocking, 1997; Finoli et al., 2001; Erdogan et al., 2003; Erdogan and Sert, 2004; Kure et al., 2004; Hayaloglu and Kirbag, 2007). One source can be raw milk (Frevel et al., 1985; Engel, 1986). Penicillium roqueforti has caused a particular problem with spoilage of packaged rye bread in Europe (see Pitt and Hocking, 1997), due to preservative resistance mentioned above, but P. paneum may also be responsible for some of this spoilage. P. roqueforti is less frequently isolated from cereals than some other species in this subgenus, but has been reported from barley, rice, flour, baked goods and refrigerated dough products (see Pitt and Hocking, 1997). Frequent reports of P. roqueforti from meats and meat products (dried beef, salami and

7.7

Genus Penicillium

257

cured meats, see Pitt and Hocking, 1997) are probably actually P. carneum. P. roqueforti has also been reported from peanuts, pecans, hazel nuts and walnuts, almonds and dried peas and fresh vegetables (see Pitt and Hocking, 1997). P. roqueforti is uncommon in the tropics: we isolated it at low levels in peanuts from the Philippines and Indonesia (Pitt et al., 1998a and our unpublished data). Additional species. Penicillium carneum (Frisvad) Frisvad is almost indistinguishable from P. roqueforti using morphological criteria (see Taxonomy). P. carneum is associated with spoilage of meat products, but has also been isolated from silage, rye bread, water, beer, cheese, mouldy bakers yeast and cork (Frisvad and Samson, 2004). P. carneum produces patulin, penicillic acid (Frisvad and Filtenborg, 1989), penitrem A, mycophenolic acid and roquefortines (Frisvad and Samson, 2004; Nielsen et al., 2006). An isolate identified as P. crustosum involved in mycotoxicosis of a man who drank mouldy beer (Cole et al., 1983) was reidentified by Frisvad and Samson (2004) as P. carneum. Additional species. Penicillium paneum Frisvad is very similar to P. carneum (see Taxonomy). It has been reported from mouldy rye bread, baker’s yeast, silage and cassava chips (Frisvad and Samson, 2004). P. paneum produces patulin and roquefortine C (Frisvad and Samson, 2004; Nielsen et al., 2006). References. Penicillium roqueforti: Samson et al. (1976); Pitt (1979b, 2000); Moreau (1980); Frisvad and Samson (2004). P. carneum and P. paneum: Boysen et al. (1996); Frisvad and Samson (2004); Karlshøj and Larsen (2005).

Penicillium solitum Westling

Fig. 7.47

Penicllium verrucosum var. melanochlorum Samson et al. Penicillium melanochlorum (Samson et al.) Frisvad

Colonies on CYA 22–28 mm diam, usually lightly radially sulcate, less commonly plicate or plane; low to moderately deep, very dense, with surface texture velutinous or less commonly granular or fasciculate; mycelium white, usually visible only at the margins; conidia usually produced abundantly, dark bluish green or dark green (25-26E-F4-7); exudate usually absent or inconspicuous, occasionally abundant or even dominating colony appearance, clear; reverse

usually pale or uncoloured, uncommonly yellow or brown, or light salmon to orange as on MEA. Colonies on MEA 20–28 mm diam, plane, low with velutinous to distinctly granular texture, or less commonly floccose; mycelium white or occasionally yellow, usually inconspicuous, but uncommonly visible over much of the colony surface; conidia abundant in most isolates, coloured as on CYA, rarely confined to central areas or absent; reverse characteristically greyish orange to brownish orange (5-6BC4-6). Colonies on G25N usually 16–22 mm diam, closely resembling colonies on CYA; conidia dark green; reverse uncoloured to pale yellow. At 58C, microcolonies or colonies up to 4 mm diam formed. No growth at 378C. On CSN, colonies 18–22 mm diam, growth strong, though not rapid; medium reaction usually alkaline (violet), but sometimes neutral or weakly acid (yellow); reverse alkaline. Conidiophores borne singly or in definite fascicles, usually from subsurface hyphae, stipes commonly 150–250 mm long, but of indeterminate length when in fascicles, with walls smooth to finely roughened or, rarely, rough; penicilli predominantly terverticillate, but appreciable numbers biverticillate or quaterverticillate in some isolates; rami usually borne singly, sometimes rough walled; phialides ampulliform but tending to acerose or cylindroidal in some isolates, commonly 9–10 mm long, with short collula; conidia spherical to subspheroidal, uncommonly broadly ellipsoidal, 3.0–4.0(–4.5) mm diam or in length, with walls smooth to very finely roughened, borne in disordered chains. Distinctive features. Conidial colour, dark bluish green to dark green, is the most useful feature distinguishing Penicillium solitum from other closely related species. On MEA, greyish to brownish orange reverse colours are a distinctive characteristic. Conidia are quite large. Growth on CSN is strong, though slow, and the colony reverse is alkaline. P. solitum can also be distinguished from P. commune by lack of reaction with the Ehrlich test (Lund, 1995a) and patterns of secondary metabolites (Lund, 1995b). Taxonomy. Accepted by Raper and Thom (1949) as a poorly defined, floccose species, Penicillium solitum was ignored for nearly 40 years. Samson et al. (1976) included it as a synonym of P. verrucosum var. cyclopium, while Pitt (1979b) considered it to be a

258

7

Penicillium and Related Genera

Fig. 7.47 Penicillium solitum (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) conidia, bar = 5 mm

synonym of P. aurantiogriseum. Cruickshank and Pitt (1987) revived P. solitum on the basis of distinctive isoenzyme patterns and morphological examination of a number of fresh isolates. Complete descriptions were provided by Pitt (2000) and Pitt et al. (1991). Physiology. Lack of recognition of this species until recently has meant that little information exists on its physiology. However, it is a typical member of Penicillium subgenus Penicillium in

showing ability to grow at low temperatures and aw and absence of growth at 378C. Mycotoxins. This species does not produce mycotoxins (Frisvad and Samson, 2004). Ecology. Penicillium solitum is a significant pathogen of pomaceous fruit (Frisvad, 1981; Pitt et al., 1991; Sanderson and Spotts, 1995; Amiri and Bompeix, 2005). It is resistant to the fungicides used to control growth of P. expansum and so its role in

7.7

Genus Penicillium

259

apple spoilage has increased (Pitt et al., 1991; Sanderson and Spotts, 1995). This species also causes spoilage of cheese (Hocking and Faedo, 1992; Lund et al., 1995; Kure et al., 2004) and has been reported from European sausages during manufacture (Andersen, 1995; Papagianni et al., 2007). Absence of other literature reports probably reflects lack of recognition, not rarity in foods. We isolated it, at a low frequency, from Thai cashews (Pitt et al., 1993), Indonesian peanuts (Pitt et al., 1998a) and Philippine peanuts, maize and mung beans (our unpublished data). References. Pitt et al. (1991); Pitt (2000); Frisvad and Samson (2004).

Penicillium verrucosum Dierckx

Fig. 7.48

Colonies on CYA 15–25 mm diam, usually closely sulcate, varying from low and velutinous to deep and fasciculate or floccose; mycelium white; conidial formation light to moderate, greyish green to dull green (26-27D-E4-5); clear to pale yellow exudate produced, copiously by some isolates; reverse yellow brown to deep brown. Colonies on MEA 12–15(–20) mm diam, sulcate, dense and velutinous or centrally floccose; mycelium white; conidial production moderate, coloured as on CYA; clear exudate occasionally produced; reverse dull brown or olive. Colonies on G25N 16–20 mm diam, plane or more commonly sulcate, velutinous to somewhat floccose; mycelium white; reverse pale to yellow. At 58C, microcolony formation at least; often colonies of 2–4 mm diam produced. No growth at 378C. On CSN, colonies usually 10–15 mm diam, growth weak, mostly subsurface; reaction neutral in medium and reverse. Conidiophores borne from subsurface or surface hyphae, stipes robust, 200–500 mm long, with walls finely to conspicuously roughened, bearing terminal penicilli; penicilli variable, some isolates producing compact terverticillate and quaterverticillate forms almost exclusively, others predominantly terverticillate and biverticillate, often with elements irregularly disposed; rami 1–2 per stipe, sometimes rough walled; phialides ampulliform, 7–9 mm long, narrowing abruptly to short collula; conidia usually spherical, 2.5–3.0 mm diam, less commonly subspheroidal to ellipsoidal, 3.0–3.5 mm long, with smooth walls, borne in disordered chains.

Distinctive features. Penicillium verrucosum is characterised by slow growth on CYA and especially on MEA, by conidia coloured relatively bright green and by the absence of other conspicuous pigmentation. Growth on CSN is weak, with no change in medium colour. P. verrucosum produces a red brown to terracotta reverse on YES agar (Frisvad and Samson, 2004). Taxonomy. Raper and Thom (1949: 486) placed Penicillium verrucosum ‘‘in the series with’’ P. viridicatum, while neither recognising the species nor placing it in synonymy. Samson et al. (1976) considered P. viridicatum and P. verrucosum as synonyms, under the name P. verrucosum var. verrucosum. Frisvad and Samson (2004) placed this species in Section Viridcata, series Verrucosa, along with P. nordicum (see Additional species, below), while placing P. viridicatum in a separate series, Viridicata. Physiology. Penicillium verrucosum grows from 0 to 318C, with the optimum at 208C. The minimum aw for germination and growth is ca. 0.80 (Northolt et al., 1979; Cairns-Fuller et al., 2005; Pardo et al., 2006c). Growth and ochratoxin A production were significantly inhibited by 50% CO2 at 0.90–0.995 aw at 258C (Cairns-Fuller et al., 2005). Growth occurs over the pH range 2.1–10.0 at least (Wheeler et al., 1991). Mycotoxins. Penicillium verrucosum is the major source of ochratoxin A in cool temperate zone commodities, especially cereals. Ochratoxin A is also produced by several Aspergillus species, notably A. carbonarius, A. westerdijkiae and A. steynii. Those species are responsible for ochratoxin A production in warmer climates. Penicillium verrucosum is endemic in cool temperate cereals, so it commonly occurs in foods for human consumption, especially breads and other cereal products. It also occurs in animal feeds and in animal tissue. Relatively high amounts are found in kidney and liver in comparison with muscle or fat. In high doses, it also appeared in eggs (JECFA, 2001). Ochratoxin A is a chronic nephrotoxin, affecting kidney function in all animal species tested, causing damage to a variety of kidney tissues. Ochratoxin A also has immunosuppressive, teratogenic and genotoxic effects in animals and possibly humans. For reviews, see Bayman and Baker (2006) and PfohlLeszkowicz and Manderville (2007). Ochratoxin A is readily absorbed through the intestines, and once it enters the blood stream, has a long half life, up to 3

260

7

Penicillium and Related Genera

Fig. 7.48 Penicillium verrucosum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

weeks in monkeys. The result of this is that in areas where ochratoxin A is frequently part of the diet, the blood of healthy humans contains detectable amounts of ochratoxin A (JECFA, 2001). Because P. verrucosum has a restricted distribution, this phenomenon is confined almost entirely to Europe. Recent indications of ochratoxin A in human blood from other parts of the world is almost certainly due to the presence of ochratoxigenic Aspergillus species in foods. Ochratoxin A also has carcinogenic properties, but the mechanism of carcinogenicity remains

unknown. The carcinogenic effects in animals are considered to be of less importance than the nephrotoxicity (JECFA, 2001). Although ochratoxin A is demonstrably toxin to animals of all kinds, its effects in humans remain unclear (JECFA, 2001). The International Agency for Research on Cancer has classified ochratoxin A as a possible human carcinogen (Group 2B), based on sufficient evidence of carcinogenicity in experimental animal studies and inadequate evidence in humans. The target organ of toxicity in all mammalian species tested is the kidneys, in

7.7

Genus Penicillium

which lesions can be produced by both acute and chronic exposure (IARC, 2002). The majority of P. verrucosum isolates are ochratoxin A producers. Some P. verrucosum isolates also produce citrinin (Frisvad and Samson, 2004). Ochratoxin A was once considered to be a possible cause of Balkan endemic nephropathy, a kidney disease with a high mortality rate in certain areas of Bulgaria, Yugoslavia and Romania. However, conclusive evidence of ochratoxin A involvement is lacking. It now seems likely that other toxins are responsible for this syndrome. Ochratoxin A is produced over the whole temperature range for growth of Penicillium verrucosum (0–318C) and down to 0.86 aw (Northolt et al., 1979). The optimum conditions on a bread dough were reported to be 0.92 aw at pH 5.6 (Patterson and Damoglou, 1986). Higher amounts of ochratoxin A were produced on wheat than other substrates including maize, peanuts, rapeseed and soybeans. Citrinin production was supported by both wheat and maize (Madhyastha et al., 1990). High concentrations of ochratoxin A were detected in exudate of P. verrucosum colonies on CYA (Gareis and Gareis, 2007). Ochratoxin A has been found in spoiled cheeses, at up to 7 mg/kg (Jarvis, 1983), and mouldy cheesecake, at up to 1 mg/kg (Piskorska-Pliszczynska and Borkowska-Opacka, 1984). A study by Biffi et al. (2004) on the ochratoxin A levels in a range of cereal products, including commercial flours and other bakery products, revealed that although all samples analysed contained ochratoxin A, most were below the legal limits set by the Italian Ministry of Health of 3 mg/kg for cereal derivatives and 0.5 mg/kg for baby foods. In another investigation of rice and rice products in Europe, Gonza´lez et al. (2006) similarly found low levels of ochratoxin A in rice and other products from both organic and non-organic sources. Ecology. Penicillium verrucosum is endemic in European and Canadian wheat, barley, rye and oats (Pohland et al., 1992; Lund and Frisvad, 2003; Samson and Frisvad, 2004; Lugauskas et al., 2006). It is a cold climate fungus, and is rare outside cool temperate zones, and away from cereals. It has also been isolated from cheese in Europe (Lund and Frisvad, 2003). References. Pitt (1987, 2000; Frisvad and Samson, 2004).

261

Additional species. Penicllium nordicum Dragoni and Canatoni ex Ramirez is very similar in appearance to P. verrucosum but its ecology is quite different. P. nordicum is generally associated with high protein substrates such as meat products (Frisvad and Samson, 2004). This species (including reports as P. viridicatum or P. verrucosum) has been isolated quite frequently from meat products in Europe (Spotti et al., 1989, 1994; Andersen, 1995; Battilani et al., 2007), from Australian salami and prosciutto in our laboratory and from cheese (Hocking and Faedo, 1992; Lund et al., 1995). P. nordicum produces a cream yellow reverse on YES medium, in contrast to the red brown reverse produced by P. verrucosum (Frisvad and Samson, 2004). All strains of P. nordicum produce ochratoxin A (Frisvad and Samson, 2004), and this toxin can be produced in meat products (Spotti et al., 2001; Pietri et al., 2006; and our unpublished observations) (Fig. 7.49).

Penicillium viridicatum Westling

Fig. 7.49

Penicillium olivinoviride Biourge Penicillium aurantiogriseum var. viridicatum (Westling) Frisvad & Filtenborg

Colonies on CYA 28–32 mm diam, radially sulcate, dense, typically relatively low, velutinous, granular or less commonly floccose; mycelium usually inconspicuous, white; conidial production moderate to heavy, yellow green, usually near cactus green (27-29E4-5), but sometimes brighter (27D4-6) or greyer (26-27D-E2-3); exudate present, clear, pale yellow, pale brown or pale pink; soluble pigment sometimes produced, orange or reddish brown; reverse in shades of orange brown, from pale, near orange white (6A2) to bright or deep, near brownish orange (6B-D7). Colonies on MEA 25–30 mm diam, plane or occasionally radially sulcate, velutinous to granular or centrally floccose; mycelium white or brown; conidial formation moderate, in yellow green colours similar to those on CYA; reverse orange brown to yellow brown. Colonies on G25N 18–22 mm diam, plane or finely radially sulcate, dense, surface granular; conidial production blue grey, yellow green or brown; reverse pale, yellow or orange. At 58C, macroscopic colonies formed, usually 2–5 mm diam. No growth at 378C.

262

7

Penicillium and Related Genera

Fig. 7.49 Penicillium viridicatum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

On CSN, colonies usually 15–22 mm diam, growth moderate to strong; medium and colony reverse response usually acid (yellow), the reverse sometimes brown also. Conidiophores borne from surface or subsurface hyphae, stipes commonly 200–300 mm long, usually with walls roughened, conspicuously so on MEA, bearing large, appressed penicilli, usually terverticillate but occasionally quaterverticillate; phialides ampulliform, 7–8(–10) mm long, usually with short collula; conidia subspheroidal to ellipsoidal, 3.0–4.0 mm long, with smooth or finely roughened walls, borne in disordered chains.

Distinctive features. On CYA, Penicillium viridicatum produces dense and compact colonies which grow moderately slowly; conidia are yellow green; stipes are rough walled. Unlike P. verrucosum, P. viridicatum grows moderately well on CSN, with an acid reaction in both medium and reverse. Like P. verrucosum, this species produces a distinctive violet brown reverse on DRYS (Frisvad, 1983) but a bright yellow reverse on YES, compared with red brown for P. verrucosum (Frisvad and Samson, 2004). Taxonomy. Penicillium viridicatum was confused with P. verrucosum for a long time (Raper and Thom, 1949; Samson et al., 1976). Ciegler et al. (1973, 1981)

7.7

Genus Penicillium

reported that P. viridicatum isolates produced two distinct classes of mycotoxins. The situation was clarified by Pitt (1987) who showed that anthraquinone toxins alone were produced by P. viridicatum. Frisvad and Filtenborg (1989) considered P. viridicatum a variety of P. aurantiogriseum, but these are separate species (Pitt and Samson, 1993; Frisvad and Samson, 2004). Physiology. A psychrophile capable of growth down to at least –28C, Penicillium viridicatum grows optimally near 238C and has its maximum at 368C (Mislivec and Tuite, 1970b). Its minimum aw for growth is 0.80–0.81 at 23–258C (Mislivec and Tuite, 1970b; Hocking and Pitt, 1979). Mycotoxins. Penicillium viridicatum produces several naphthoquinones, notably xanthomegnin and viomellein capable of causing liver and kidney damage in mice, rats and pigs. Both compounds caused illness in young mice when administered at 450 mg/kg body weight (Carlton et al., 1973, 1976). Viomellein has been found in nature, in a Danish barley sample, at a level of 1 mg/kg (Hald et al., 1983). This level appears to be too low to be considered hazardous. Eleven of 19 samples of poor quality cereals and compounded feeds examined in the United Kingdom contained xanthomegnin and viomellein and a third related compound, vioxanthin. Levels of each up to 2 mg/kg were found, again a significant finding of naturally occurring mycotoxins, but below a clearly hazardous level (Scudamore et al., 1986). The role, if any, of these toxins in natural disease of animals remains unknown. Ecology. The reported association of Penicillium viridicatum with European barley (Krogh et al., 1973) and Canadian wheat (Wallace et al., 1976) is incorrect: P. verrucosum is the species involved. P. viridicatum is probably rather rare, however, its occurrence in Canadian grain has been verified (Mills et al., 1995), and Frisvad and Samson (2004) report P. viridicatum from maize, wheat, barley, beans and peas from Europe, the United States, Canada, Ethiopia and Taiwan. Penicillium viridicatum has been reported as weakly pathogenic, causing spoilage of grapes and melons in storage. This species has also been reported frequently from meats, dried fruit, spices, pasta, health foods, peanuts, pistachio nuts, almonds (see Pitt and Hocking, 1997) and cheese (Hayaloglu and Kirbag, 2007). We isolated P. viridicatum from Philippine maize, peanuts, milled rice, soybeans and black pepper and dehydrated infant food (our unpublished data).

263

References. Pitt (1979b, 2000); Samson et al. (1976) and Domsch et al. (1980), both under the name Penicillium verrucosum var. verrucosum; Frisvad and Samson (2004).

7.7.4 Penicillium subgenus Biverticillium Dierckx (Penicillium Sect. Biverticillata-Symmetrica Thom) In Penicillium subgenus Biverticillium, penicilli are characteristically biverticillate, though with sometimes a proportion terverticillate, and always terminal; metulae are numerous, in symmetrical appressed or divergent verticils, and of approximately equal length to phialides; phialides are typically acerose (shaped like a pine needle), with conical collula, tapering to narrow apical pores; conidia in species considered here are ellipsoidal to fusiform. Colonies on CYA at 258C commonly show yellow or red colours in mycelium, exudate, soluble pigment and/ or reverse. Growth at 378C commonly occurs. Growth at 58C is confined to a single species considered here, Penicillium rugulosum. Growth on G25N is slow; colonies are not more than 10 mm diam in 7 days at 258C. Subgenus Biverticillium comprises a well circumscribed group of species with many characters in common, of which penicillus structure, phialide shape, colony pigmentation and growth rate on G25N are especially significant. The most readily recognised feature of isolates from this subgenus is the orderly, characteristically biverticillate penicillus. Only a few species in subgenus Furcatum produce penicilli which are superficially similar: however, such species grow more than 10 mm in 7 days on G25N and produce metulae which are consistently longer than phialides. Where species in subgenus Biverticillium produce teleomorphs, they belong in Talaromyces. Recent molecular studies have shown that some species in subgenus Biverticillium have Talaromyces species as their nearest neighbours, indicating multiple losses of the teleomorph. In particular, T. wortmannii was shown to be very closely related to Penicillium variabile, and T. flavus with P. minioluteum and P. funiculosum (LoBuglio et al., 1993; Pitt, 1995). Most species in subgenus Biverticillium are uncommon in foods. All species in this subgenus

264

7

have relatively high water requirements and appear to be associated primarily with soil or moist, decaying vegetation. Perhaps as a consequence, the six

Penicillium and Related Genera

species considered here are all well known as biodeteriorative agents in situations where moisture is not a factor limiting growth.

Key to Penicillium subgenus Biverticillium species included here 1

Colonies on MEA exceeding 25 mm diam Colonies on MEA not exceeding 25 mm diam

2 (1)

Red soluble pigment produced by colonies on CYA Soluble pigment on CYA pink or absent

3 (2)

Mycelium on CYA and MEA bright yellow; stipes more than 100 mm long Mycelium on CYA and MEA not bright yellow; stipes less than 100 mm long

4 (1)

Colonies on CYA deep and convex; the dominant colony colour bright yellow or orange mycelium Colonies on CYA low and velutinous; the dominant colony colour green conidia

5 (4)

2 4

Colonies on CYA at 258C not exceeding 12 mm diam; sometimes germination at 58C Colonies on CYA at 258C exceeding 12 mm diam; no germination at 58C

Penicillium funiculosum Thom

Fig. 7.50

Colonies on CYA 25–40 mm diam, plane, usually 2–5 mm deep, with conspicuous ropes of aerial hyphae (funicles), occasionally almost velutinous; mycelium salmon to peach (6-7A4-5) or brownish orange to brownish red (7-8C5-6); conidia moderately abundant, dull green (26-27C-D3-4); clear exudate and pink soluble pigment produced by some isolates; reverse pale, brown, or more commonly deeply pigmented, brownish red to red (810B-C8). Colonies on MEA 25–45 mm diam, usually conspicuously funiculose; mycelium usually white, less commonly salmon (6A4) to brownish red (8-9C6-7); conidial production moderate to heavy, coloured as on CYA; reverse pale, brown or orange to reddish brown. Colonies on G25N 3–8 mm diam, plane, usually funiculose; reverse pale to olive. No germination at 58C. At 378C, colonies 20–45 mm diam, usually similar to those on CYA at 258C or more floccose, with reverse pale, brown or reddish brown. Conidiophores borne from aerial hyphae, usually from well defined funicles, stipes short, commonly 25–40(–60) mm long, in uncommon isolates shorter or longer, with walls smooth to finely roughened, bearing terminal biverticillate penicilli, or occasionally more complex or irregular forms; metulae and phialides closely appressed, acerose, 9–11 mm long,

P. purpurogenum 3 P. pinophilum P. funiculosum

P. islandicum 5 P. rugulosum P. variabile

phialides with gradually tapering collula; conidia small, cylindroidal to ellipsoidal, 2.2–3.0 mm long, smooth walled, borne in short columns. Distinctive features. Colonies of Penicillium funiculosum often produce well defined ropes of hyphae (funicles) bearing short conidiophores at right angles. Growth at 378C is rapid, similar to that on CYA at 258C or more floccose; penicilli are closely appressed and conidia are pale greyish green. Physiology. Mislivec and Tuite (1970b) reported that Penicillium funiculosum grows optimally at ca. 308C, with a minimum near 88C; the maximum growth temperature is near 428C (Domsch et al., 1980). The minimum aw for germination and growth is 0.90 at 23, 25 and 308C (Mislivec and Tuite, 1970b; Hocking and Pitt, 1979). This species is very acid tolerant; we have isolated it from soils on media of pH 2.0. Mycotoxins. Penicillium funiculosum strains toxic to day old cockerels were isolated from chestnuts (Wells and Payne, 1975) and pecans (Wells, 1980). The toxic compounds have not been characterised. Ecology. Penicillium funiculosum has been found quite frequently in foods, including fruit, nuts and cereals. It is one cause of core rot in apples (Combrink et al., 1985; Vismer et al., 1996). P. funiculosum causes diseases in onions (Hussein et al., 1977) and blemishes in pineapples (Matos et al., 2005). It

7.7

Genus Penicillium

265

Fig. 7.50 Penicillium funiculosum (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) conidia, bar = 5 mm

was reported as causing spoilage in pecans, rots in kola nuts and as being common in peanuts both before and after harvest (see Pitt and Hocking, 1997). This species also has a close association with maize (see Pitt and Hocking, 1997) and maize products (Ribeiro et al., 2003). It was present in 42% of maize samples we examined from Thailand, at levels up to 56% of kernels in a sample and in 4% of all kernels examined. Levels in Philippine and Indonesian maize were lower (Pitt et al., 1993, 1998a).

Isolations of Penicillium funiculosum have also been made from walnuts, wheat and flour, pasta, barley, soybeans, rice (see Pitt and Hocking, 1997), fresh cabbages (Lugauskas et al., 2005) and parboiled rice (Jayaraman and Kalyanasundaram, 1994). Other sources include dried peas, miso, biltong, a Nigerian spice, yam chips and carbonated soft drinks (see Pitt and Hocking, 1997). As well as peanuts and maize, mentioned above, we isolated Penicillium funiculosum at low levels

266

7

from cashews and black rice in Thailand (Pitt et al., 1994). References. Pitt (1979b, 2000); Domsch et al. (1980).

Penicillium islandicum Sopp

Fig. 7.51

Colonies on CYA 17–22 mm diam, plane or centrally raised, velutinous to lightly floccose; mycelium intensely coloured, dominating the colony appearance, deep orange to brownish orange (6A-C8) or copper (7C8); conidia produced in moderate numbers, usually enveloped by the mycelium, but if conspicuous coloured greyish turquoise to greyish green (2425D4); clear to pale yellow exudate sometimes produced; reverse very strongly coloured, orange to rust brown or burnt sienna (6-7B-E8), sometimes reddish brown (9D-E8). Colonies on MEA 17–22 mm diam, similar to those on CYA but with mycelium usually less dominant; conidial production heavy, greyish turquoise (24D5); reverse commonly with central areas brown to reddish brown (7-8B-E8). Colonies on G25N 4–9 mm diam, similar in appearance and colouration to those on CYA and MEA. No germination at 58C. At 378C, colonies 10–20 mm diam, deep and floccose; mycelium white or coloured as at 258C; conidia absent to abundant, dark green near bottle green (26F5); reverse pale or as at 258C. Conidiophores borne from aerial hyphae, stipes 30–60 mm long, usually smooth walled, bearing terminal penicilli, typically biverticillate but not uncommonly bearing an appressed ramus; phialides acerose, 7–8 mm long, with abruptly narrowing collula; conidia broadly ellipsoidal to subspheroidal, mostly 3.0–3.5 mm long, with smooth heavy walls, borne in short, irregular to well defined columns. Distinctive features. Among the most readily recognised Penicillium species, P. islandicum produces compact colonies with brilliant orange to brown mycelium and reverse on CYA. Conidia are usually blue and are smooth walled. Physiology. The optimum temperature for growth of Penicillium islandicum has been reported as 318C, with a minimum and maximum of 10 and 388C, respectively (Domsch et al., 1980). However, a consistent ability to grow nearly as rapidly at 378C as at 258C indicates the maximum growth temperature may be nearer 428C. The minimum aw for growth at 318C is 0.83 (Ayerst, 1969) and at 258C,

Penicillium and Related Genera

0.86 (Hocking and Pitt, 1979). Growth occurs over the pH range 2.1–9.2 at least (Wheeler et al., 1991). Mycotoxins. Penicillium islandicum produces at least four mycotoxins, unique to the species. Cyclochlorotine and islanditoxin are chlorine containing cyclic peptides which have the same toxic moiety, a pyrrolidine ring with two attached chlorine atoms and share a number of other physical and chemical properties (Scott, 1977). Both compounds are very toxic: cyclochlorotine has an oral LD50 in mice of 6.5 mg/kg, while that of islanditoxin by subcutaneous injection was 3 mg/kg. Fed to mice at the rate of 40 mg/day, cyclochlorotine caused liver cirrhosis, fibrosis and tumours (Uraguchi et al., 1972). Luteoskyrin is a dimeric anthraquinone and erythroskyrin a heterocyclic red pigment. Both are liver and kidney toxins, though less acutely toxic than cyclochlorotine. Luteoskyrin is also carcinogenic. The toxic ‘‘yellow rice’’ syndrome, described in Japan last century, has resulted in Japanese scientists taking a particular interest in Penicillium islandicum, a species which causes yellowing of rice (Saito et al., 1971a). However, the practical importance of the toxins produced by this species remains unclear. P. islandicum has not been commonly found growing in nor causing spoilage of foods. Ecology. A marginal xerophile, Penicillium islandicum is more tolerant of low aw than most other species in subgenus Biverticillium. This species is an active agent of spoilage in cereals stored a little above safe moisture contents. It occurs in soil, though not abundantly, and perhaps also is a weak animal pathogen (Pitt, 1979b; p 447). It appears to be more widespread in tropical than temperate regions. Penicillium islandicum is often associated with rice and has been reported from Japan (Sakai et al., 2005) and South America (Tonon et al., 1997). Other reports include flour, peanuts, pecans and soybeans (see Pitt and Hocking, 1997), without mention of spoilage. We isolated P. islandicum from Southeast Asian commodities including maize from the Philippines (our unpublished data), peanuts from Indonesia and milled rice and mung beans from both countries (Pitt et al., 1998a). Levels were always low, without evidence of spoilage or toxin production. References. Pitt (1979b, 2000); Domsch et al. (1980).

7.7

Genus Penicillium

267

Fig. 7.51 Penicillium islandicum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d, e) penicilli, bars = 10 mm; (f) conidia, bar = 5 mm

Penicillium pinophilum Hedgcock

Fig. 7.52

Colonies on CYA 20–30 mm diam, plane or less commonly sulcate, texture rather loose and floccose, often with tufts of rudimentary funicles also; mycelium white at the margins and also sometimes in superficial overlays, but predominantly slightly brighter than sulphur yellow (1A5), usually produced in a relatively dense overlay over most or all of the colony area, dominating colony appearance and masking conidial colour; conidial formation moderate, usually dark green (28F5) but often appearing lighter due to the yellow mycelium;

exudate sometimes present in small amounts, clear to red; reverse usually strongly coloured, golden brown (5C-D7), sometimes masked by or mixed with deep red shades as well. Colonies on MEA 25–35 mm diam, plane, deep and rather floccose; characteristics similar to colonies on CYA, except for paler reverse colouration and absence of white mycelial overlays. Colonies on G25N 2–5 mm diam, rarely as large as 8 mm, low and sparse; mycelium white; reverse pale, brown or olive. No germination at 58C. At 378C colonies 20–40 mm diam, plane or lightly sulcate, sometimes low and velutinous and coloured yellow green, more commonly deeply

268

floccose with white mycelium predominant; conidial production absent to moderate, bluish to greenish grey; clear exudate usually present; reverse usually yellowish or reddish brown. Sclerotia produced by some isolates, dark red to reddish brown, of variable size, composed of soft compacted hyphae, not of sclerotioid tissue. Teleomorph not known. Conidiophores borne from aerial hyphae, stipes 150–180 mm long, smooth walled, often with vesiculate apices, bearing terminal biverticillate penicilli; metulae in verticils of 8–12 or even

7

Penicillium and Related Genera

16, 9–12 mm long, forming a 60–908 angle and in the latter case 12–15 mm across the apices; phialides acerose, 8–10 mm long, with gradually tapering collula; conidia subspheroidal, rarely ellipsoidal, commonly 2.5–2.8 mm long, with walls relatively heavy and smooth to finely roughened, borne in short disordered chains or loose columns. Distinctive features. Penicillium pinophilum is characterised by the production of bright yellow mycelium which dominates colony appearance on CYA and MEA. Conidia are subspheroidal to

Fig. 7.52 Penicillium pinophilum (a) colonies on CYA and MEA, 7 d, 258C; (b, c) penicilli, bars = 10 mm; (d) conidia, bar = 5 mm

7.7

Genus Penicillium

269

ellipsoidal, less than 3 mm in long axis, and with smooth to finely roughened walls. Physiology. Little is known about the physiology of this species. It grows rather strongly at 378C. From its taxonomic position, we can infer that it not a xerophile and is similar in most properties to Penicillium funiculosum. Mycotoxins. Mycotoxin production has not been reported. Ecology. Although Penicillium pinophilum has been reported from foods only occasionally, in our experience it is not rare. It was reported from fresh meat by Nassar and Ismail (1994) and it was found in low levels on Spanish grapes (Bau et al., 2005). We isolated it from Philippine peanuts, and in one sample 85% of kernels were infected: this was exceptional, however (our unpublished data). We also found it in maize from Thailand and the Philippines and peanuts from Indonesia (Pitt et al., 1993, 1998a). References. Pitt (1979b, 2000).

Penicillium purpurogenum Stoll

Fig. 7.53

Colonies on CYA 15–30 mm diam, plane or radially sulcate, dense, usually velutinous; mycelium bright yellow or red due to encrusted hyphae; conidial production moderate to heavy, dark green (2527E-F5); exudate orange to red; soluble pigment vivid red (10A8); reverse dark red or purple, approaching black. Colonies on MEA 22–35 mm diam, plane, dense, velutinous; mycelium white to bright yellow; conidia abundantly produced, dark green (26-27E-G4-7); reverse usually pale, often brown or dull red centrally. Colonies on G25N microscopic to 6 mm diam, coloured as on CYA; reverse pale to deep brown. No germination at 58C. At 378C, colonies commonly 12–22 mm diam, usually similar to those on CYA at 258C, occasionally lacking soluble red pigment. Conidiophores borne from surface or aerial mycelium, stipes 70–300 mm long, smooth walled, bearing terminal biverticillate penicilli; penicilli narrow, metulae and phialides appressed, 10–14 mm long; conidia ellipsoidal, sometimes becoming subspheroidal at maturity, 3.0–3.5 mm long, with walls smooth, finely roughened or warty, borne in short irregular chains. Distinctive features. An intense, rapidly diffusing red pigmentation on CYA at both 25 and 378C is the

most striking characteristic of Penicillium purpurogenum. In the (rare) absence of such pigmentation, other diagnostic features include conidia very dark green on both CYA and MEA; moderate growth at 378C (15–22 mm in 7 days); narrow penicilli and heavy walled ellipsoidal to subspheroidal conidia. Physiology. Minimum and optimum temperatures for growth by Penicillium purpurogenum are reported as 12 and 308C (Mislivec and Tuite, 1970b); from our growth data, a maximum growth temperature near 408C would be expected. Hocking and Pitt (1979) reported 0.84 as the minimum aw for germination and growth of this species. Like P. funiculosum, P. purpurogenum is highly tolerant of acid and can be readily isolated on media of pH 2.0 (Pitt, unpublished). Mycotoxins. The rubratoxins were originally reported from Penicillium rubrum, a species synonymised with P. purpurogenum by Pitt (1979b). Rubratoxins were originally suggested as causal agents of mouldy corn toxicosis, or haemorrhagic anaemia in chickens (Burnside et al., 1957; Forgacs et al., 1958). However, later studies (Wyatt and Hamilton, 1972) concluded that rubratoxins were not the cause of these syndromes. Recent taxonomic studies (International Commission on Penicillium and Aspergillus, unpublished) indicate that rubratoxins are produced by only three known fungal isolates which represent an undescribed species. Ecology. A marginal xerophile and a recognised biodeteriogen (Pitt, 1981), Penicillium purpurogenum has been reported as a cause of spoilage in pineapples in India, pears and Averrhoa bilimbi fruit in Sri Lanka (see Pitt and Hocking, 1997). It has been isolated from a wide variety of other foodstuffs, but seldom as a cause of spoilage: from cassava (C.J. Rabie, unpublished), maize, rice, wheat, barley, peanuts, soya beans and kidney beans (Aziz et al., 2006), pecans and betel nuts (see Pitt and Hocking, 1997). It has also been found in processed meats (see Pitt and Hocking, 1997). References. Pitt (1979b, 2000).

Penicillium rugulosum Thom

Fig. 7.54

Penicillium tardum Thom

Colonies on CYA 4–8 mm or occasionally 12 mm diam, plane, low and dense, velutinous; mycelium usually inconspicuous, mainly white but yellow or

270

7

Penicillium and Related Genera

Fig. 7.53 Penicillium purpurogenum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

red encrusted hyphae often visible under magnification; conidia abundant, greenish grey to dark green (26-27E-F2-4); exudate and soluble pigment usually absent; reverse pale, dull olive or brown. Colonies on MEA 10–20 mm diam, similar to those on CYA except for more numerous and conspicuous yellow encrusted hyphae. Colonies on G25N 2–8 mm diam, velutinous; conidial production moderate, coloured as on CYA, other pigmentation absent. At 58C, in some isolates germination of conidia, in others germination limited or absent. At 378C, typically no growth, rarely colonies up to 4 mm diam formed (Fig. 7.54).

Conidiophores borne from surface or aerial hyphae, stipes 70–100 mm long, with thin, smooth walls; penicilli basically biverticillate, but more complex structures often present, for example rami in verticils of up to 4, or rami and metulae from a common origin; metulae appressed; phialides acerose, tending towards ampulliform or less commonly cylindroidal, 8–11 mm long, typically with collula tapering to narrow pores, but sometimes untapered or even enlarging at the apices; conidia ellipsoidal, 3.0–3.5 mm long, with heavy, smooth to rough walls, borne in short to quite long, irregular columns.

7.7

Genus Penicillium

271

Fig. 7.54 Penicillium rugulosum (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d) penicilli, bars = 10 mm; (e) conidia, bar = 5 mm

Distinctive features. Penicillium rugulosum is characterised by very slow growth, velutinous colonies, and usually by germination at 58C. Although this species clearly belongs to subgenus Biverticillium, penicilli may be of atypical structure at times, and may also bear ampulliform to cylindroidal phialides. Physiology. From growth data (Pitt, 1973), this species grows between 5 and 378C; the optimum

would be expected to be near 258C. Snow (1949) reported 0.86 aw to be its minimum for germination. Mycotoxins. Production of rugulosin by Penicillium rugulosum was reported by Yamazaki et al. (1971) and El-Banna et al. (1987b). Although clearly a toxic compound when given intraperitoneally, with an LD50 of 83 mg/kg in mice (Cole and Cox, 1981), rugulosin may be only a laboratory toxin. Disease due to rugulosin has not been reported.

272

Ecology. Because of its very slow growth, Penicillium rugulosum is easily overlooked in isolation procedures. It is probably a more commonly occurring species than records would suggest, and it appears to be widely distributed. This species can be a plant pathogen: it was described by Charles Thom in 1910 from rotting potato tubers, and was isolated by Barkai-Golan (1974) from decaying cold stored apples and the surface of fresh, healthy apples (Amiri and Bompeix, 2005). P. rugulosum

7

Penicillium and Related Genera

has been reported quite frequently from dried and processed meats (see Pitt and Hocking, 1997). It has also been found in cheese (Hocking and Faedo, 1992; Lund et al., 1995), rice, maize, barley, wheat, peanuts, soybeans and kidney beans (Aziz et al., 2006), flour and pecans (see Pitt and Hocking, 1997). We isolated it at low levels from Thai cashews and Philippine soybeans (Pitt et al., 1994, 1998a). References. Pitt (1979b, 2000).

Fig. 7.55 Penicillium variabile (a) colonies on CYA and MEA, 7 d, 258C; (b, c, d, e) penicilli, bars = 10 mm; (f) conidia, bar = 5 mm

7.7

Genus Penicillium

Penicillium variabile Sopp

273

Fig. 7.55

Colonies on CYA and MEA 15–22 mm diam, plane or irregularly furrowed, low and dense, velutinous; mycelium commonly sulphur yellow (1A4-5), usually conspicuous only at the margins and near colony centres; conidia moderately to abundantly produced, greenish grey (25-27D2); clear exudate occasionally produced; reverse on CYA deep orange to raw umber (5A-F8), on MEA more muted, usually deep orange to brownish yellow (5A-C8). Colonies on G25N microscopic to 9 mm diam, velutinous, sometimes heavily sporing; other colouration absent. No germination at 58C. At 378C, response variable, commonly no growth but sometimes colonies up to 5 mm diam produced. Conidiophores borne mostly from surface hyphae, stipes commonly 100–200 mm long, but if from aerial growth much shorter, 10–30 mm long, smooth walled; penicilli usually biverticillate, sometimes with a subterminal ramus or with concurrent metulae and phialides, 8–12 mm long, appressed to quite divergent; conidia narrowly ellipsoidal, 3.0–4.0(–6.0) mm long, with walls smooth or faintly roughened, borne in closely packed but disordered chains. Distinctive features. Penicillium variabile produces strictly velutinous colonies with abundant greenish grey conidia and with sulphur yellow

mycelium conspicuous only in peripheral and central areas. Growth is poor to negative at 378C. Unlike P. rugulosum, penicilli of P. variabile are typical of subgenus Biverticillium and conidia never germinate in 7 days at 58C. Physiology. Mislivec and Tuite (1970b) reported 12 and 308C to be the minimum and optimal growth temperatures for Penicillium variabile; the maximum is near 378C (Pitt, 1973). The minimum aw for germination and growth is 0.86 (Hocking and Pitt, 1979). Mycotoxins. Like Penicillium rugulosum, P. variabile produces rugulosin (Yamazaki et al., 1972; El-Banna et al., 1987b). This is probably not an important toxin, and there appear to be no records of P. variabile causing disease. Ecology. Although the primary habitats of Penicillium variabile, like other species in this subgenus, would be expected to be soil and decaying vegetation, it has quite commonly been found in foods. Isolations have come principally from cereals: wheat and flour, maize, rice and barley (see Pitt and Hocking, 1997). Some other reported sources have been processed meats (Cantoni et al., 2007), peanuts (Pitt et al., 1998a), biltong, cheese, pecans, walnuts and betel nuts (see Pitt and Hocking, 1997). References. Pitt (1979b, 2000); Domsch et al. (1980).

“This page left intentionally blank.”

Chapter 8

Aspergillus and Related Teleomorphs

Thriving in, or at least tolerant of, elevated temperatures and reduced water activities, species of Aspergillus and its teleomorph Eurotium are the epitome of spoilage fungi. There are few kinds of foods, commodities and raw materials from which Aspergilli cannot be isolated consistently. Aspergillus species compete with Penicillium and Fusarium species for dominance among the world’s fungal flora. Aspergillus lacks the sheer numbers and diversity of Penicillium species but compensates by the ability to grow at higher temperatures or lower water activities or both. Aspergilli usually grow more rapidly than Penicillia, although they take longer to sporulate, and generally produce spores which are more resistant to light and chemicals. It is fair to say that Aspergillus species dominate spoilage in the tropics in the way Penicillium species do in temperate zones. A small number of Aspergillus species are associated with plants, and here Aspergillus is more

directly in competition with Fusarium than with Penicillium. Aspergillus is a genus of Hyphomycetes characterised, in general terms, by the formation of conidiophores with large, heavy walled stipes and swollen apices, termed vesicles. Vesicles are usually roughly spherical, but are elongated or less conspicuously swollen in a few species. Vesicles bear crowded phialides, or metulae and phialides, which are characteristically all borne simultaneously (Fig. 8.1a). This character unequivocally distinguishes Aspergillus from Penicillium and the other genera grouped with it in Chapter 7. Phialide production in Penicillium and related genera is always successive, not simultaneous (Fig. 8.1b). Ready differentiation of these genera may be obtained by microscopic examination of developing conidiophores picked from near the colony margins. The presence of immature metulae or phialides all at the same stage of

Fig. 8.1 Scanning electron micrographs: (a) Aspergillus oryzae showing simultaneous production of phialides, bar = 50 mm; (b) Penicillium janczewskii showing successive production of phialides, bar = 10 mm

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_8, Ó Springer ScienceþBusiness Media, LLC 2009

275

276

development indicates Aspergillus; structures with some phialides producing conidia while one or more others are still developing indicates Penicillium or a related genus. Two other useful features are characteristic of most, but not all, Aspergillus species. First, stipes are usually formed from a short cell termed a footcell within a fertile hypha. Second, stipes are usually nonseptate, so that the vesicle, stipe and footcell all form a very large single cell. Penicillium stipes are usually septate and footcells are exceptional. The authoritative text on Aspergillus for the past 40 years has been ‘‘The Genus Aspergillus’’ by Raper and Fennell (1965), and this monograph deserves its status. However, in using that monograph, the reader must be aware of the ways in which that classification is considered deficient or unacceptable in current systematics. First, Raper and Fennell (1965) declined to give status to teleomorph names, an approach with some logic and merit (Raper, 1957), but which contravenes the Botanical Code. Never accepted by nomenclaturalists, that approach has lost favour among industrial mycologists as well. Second, Raper and Fennell (1965) referred to metulae and phialides as ‘‘primary and secondary sterigmata’’. The term sterigma has rightly been confined to Basidiomycete terminology in recent years (Kendrick, 1971). Third, subdivision of the genus was based on species ‘‘groups’’, i.e. species related to Aspergillus ochraceus were referred to as belonging to the ‘‘Aspergillus ochraceus group’’. The use of the term ‘‘group’’ in this way has no status under the Botanical Code and has been sharply criticised (Benjamin, 1966). Fourth, Raper and Fennell (1965) often failed to observe the rules of priority in accepting particular species names and, fifth, they failed to typify species. The failure to typify names by Thom and Raper and their coworkers over the years led to concept changes for some species, e.g. Eurotium amstelodami and A. penicillioides. For a discussion of these problems see Pitt (1989a) and Pitt and Samson (1990a). The systematic difficulties arising from Raper and Fennell (1965) have been addressed by modern taxonomists. Subramanian (1971) and Malloch and Cain (1972) provided names for Aspergillus teleomorphs. Gams et al. (1985) developed a new classification of Aspergillus based on subgenera and series to replace the incorrect term ‘‘group’’. Gams and Samson (1985) provided typifications of Aspergillus and its associated teleomorphs, while Samson and

8 Aspergillus and Related Teleomorphs

Gams (1985) typified the species of Aspergillus accepted by Raper and Fennell (1965). Samson (1979) produced a useful compendium and taxonomic outline of the 90 species described between 1965 and that date. A guide to common Aspergillus species and Aspergillus teleomorphs which incorporated all of these advances was published by Klich (2002). Pitt and Samson (1993) produced a list of Names in Current Use (NCU) for the family Trichocomaceae, which includes Aspergillus and related teleomorphs. By this means the names used by Raper and Fennell (1965) were given de facto protection against earlier valid names. This was especially significant in Aspergillus, where a number of common names cannot be justified under the Botanical Code. One particularly important name, A. niger, was conserved under the provisions of the Botanical Code (Frisvad et al., 1990c; Kozakiewicz et al., 1992) because this name was known to be predated by A. phoenicis (Corda) Thom and A. ficuum (Reichardt) Hennings. The names used here are in all cases those given protected status by the NCU list. Teleomorphs. As is the case with Penicillium and Fusarium, most Aspergillus species do not produce any known teleomorph. Those that do all form cleistothecia or reduced stromata resembling cleistothecia and are classified in eight Ascomycete genera. Three are significant in foods: Eurotium, Neosartorya and Emericella. Each of these genera is discussed below, in alphabetical order, before the strictly anamorphic species of Aspergillus. Rather than providing a separate key, diagnoses of the distinctions between these three genera are given below and then keyed in the general Aspergillus key. Eurotium species produce bright yellow cleistothecia and pale yellow ascospores. Heads producing conidia are formed from phialides only. All species are xerophilic and grow more vigorously on G25N than on CYA or MEA. If a Eurotium isolate is entered in the key in Chapter 5 it will usually be recognised as a Eurotium in the general xerophile key in Chapter 9. Neosartorya species also produce heads formed from phialides alone, but in this case cleistothecia are white and ascospores are uncoloured. Colonies grow rapidly on CYA and MEA at 25 and 378C. Species are not xerophilic. Emericella species produce heads with both metulae and phialides and white cleistothecia producing red or purple ascospores. The cleistothecia are surrounded by Hu¨lle cells, which are thick walled roughly spherical

8

Aspergillus and Related Teleomorphs

cells resembling chlamydoconidia. Growth patterns in Emericella species are similar to those of Neosartorya. In Emericella, conidiophores of the Aspergillus anamorph are usually abundant, in Neosartorya they are often only visible through the stereomicroscope, while in Eurotium occurrence is variable. In all three genera the conidial colour is grey green. Classification. Classification of Aspergillus into subgenera and sections relies primarily on four features: the presence of a teleomorph and its characteristics, the presence or absence of metulae, the arrangement of metulae or phialides on the vesicle, and colony colours. In species lacking teleomorphs, Aspergillus colony colours are dominated by conidial colour. These colours are consistently associated with particular species.

277

The subgeneric and sectional classification of Gams et al. (1985) adopted by Klich (2002) has been followed here. Colony diameters are a valuable additional character. In keeping with standardisation of media and growth conditions throughout this book, colony diameters at 58C are incorporated in the descriptions which follow. However, it should be noted that very few Aspergillus or Aspergillus teleomorph species germinate at 58C in 7 days, so if the user knows in advance that an isolate is an Aspergillus or related teleomorph, the 58C plate is logically omitted. CY20S, the medium used here for Eurotium identification, can also be a useful identification medium for most Aspergillus species, but has not been included here. However, Table 8.1 provides figures for colony diameters on CY20S which may be of value to some users.

Key to Aspergillus species and teleomorphs included here 1

Colonies on CYA at 25 and 378C both exceeding 35 mm diam Colonies on CYA at 25 or 378C not exceeding 35 mm diam

2 (1)

Colonies black or grey Colonies white or coloured

3 (2)

Colonies black, exceeding 50 mm diam at 258C Colonies grey, not exceeding 50 mm diam at 258C

4 (3)

Vesicles bearing metulae and phialides Vesicles bearing phialides alone

5 (4)

Conidia less than 6 mm diam Conidia more than 6 mm diam

6 (2)

Colonies white Colonies coloured

7 (6)

Colonies blue Colonies yellow, green or brown

8 (7)

Conidia dark green; developing cleistothecia present, surrounded by cells like chlamydoconidia (Hu¨lle cells) Conidia yellow, yellow green or brown; developing cleistothecia not present

9 (8)

Conidia yellow green or yellow Conidia brown or olive brown

10 (9)

Conidia with relatively thin walls, smooth or finely roughened, spherical to broadly ellipsoidal; vesicles up to 50 mm diam, usually metulae present

Conidia with consistently rough, thick walls, spherical; vesicles not usually exceeding 30 mm diam; usually only a low proportion of heads with metulae

2 12 3 6 4 A. ustus 5 A. aculeatus A. japonicus A. niger A. carbonarius Genus Neosartorya 7 A. fumigatus 8

Genus Emericella 9 10 11

A. flavus A. oryzae A. nomius A. parasiticus

278

8 Aspergillus and Related Teleomorphs

Key to Aspergillus species and teleomorphs included here (continued) 11 (9)

12 (1)

Colonies olive brown on CYA and MEA at 258C; conidia 5–8 mm diam, rough walled; heads radiate Colonies brown on all media; conidia less than 3 mm diam, smooth walled; heads developing into long columns

A. tamarii A. terreus A. ustus

Colonies grey Colonies white or brightly coloured

13

13 (12)

Colonies white, off-white or cream Colonies coloured

14 15

14 (13)

Vesicles fertile over the entire area; metulae more than 10 mm long Vesicles fertile over the upper half to two-thirds; metulae less than 10 mm long

15 (13)

Developing yellow cleistothecia present, particularly in colonies on G25N Developing cleistothecia not present

16 (15)

Conidia in shades of yellow, orange or brown Conidia green or blue

17 (16)

Colonies on CYA not exceeding 40 mm diam; conidia in yellow or orange shades Colonies on CYA exceeding 40 mm diam; conidia pale brown

A. candidus A. niveus

Genus Eurotium 16 17 19 18 A. ochraceus A. westerdijkiae A. steynii

18 (17)

Colonies greyish orange, conidia 2–3 mm diam, smooth walled Colonies greyish yellow to olive, conidia 4–5 mm diam, with smooth or rough walls

A. flavipes A. wentii

19 (16)

Colonies on CYA exceeding 30 mm diam; vesicles more than 50 mm diam Colonies on CYA not exceeding 30 mm diam; vesicles less than 25 mm diam

A. clavatus

20

20 (19)

Colonies on CYA exceeding 15 mm diam; heads with metulae Colonies on CYA not exceeding 15 mm diam; heads with phialides only

21 22

21 (20)

Conidia green Conidia blue

22 (20)

Colonies on CYA and MEA 6 mm or more diam; conidia cylindrical to barrel shaped, borne in columns Colonies on CYA and MEA not exceeding 6 mm diam; conidia subspheroidal to ellipsoidal, borne in radiate heads

A. versicolor A. sydowii

A. restrictus A. penicillioides

Table 8.1 Colony diameters expected for Aspergillus species and teleomorphs Species Emericella nidulans Eurotium amstelodami E. chevalieri E. halophilicum E. herbariorum E. repens E. rubrum Neosartorya fischeri Aspergillus aculeatus

CYA 40–50 14–18 16–25 0 0–10 10–20 10–20 50–70 65–70

MEA 35–60 16–25 16–25 0 0–10 10–20 5–20 60–70 45–70

G25N 10–15 25–35 20–30 0–2 25–35 25–45 30–45 8–12 15–20

378C 50–70 0–15 0–10 0 0 0 0 60–70 5–30

CY20S 40–50 30–45 45–65 0 25–35 45–65 35–60 60–70 65–70

8.1

Genus Emericella

279

Table 8.1 Colony diameters expected for Aspergillus species and teleomorphs (continued) A. candidus A. carbonarius A. clavatus A. flavipes A. flavus A. fumigatus A. japonicus A. niger A. niveus A. nomius A. ochraceus A. oryzae A. parasiticus A. penicillioides A. restrictus A. steynii A. sydowii A. tamarii A. terreus A. ustus A. versicolor A. wentii A. westerdijkiae

15–20 60–70 30–45 15–30 60–70 40–60 45–70 60–70 18–30 60–70 40–45 55–70 50–70 0–5 6–12 33–45 18–30 50–65 40–50 30–40 15–25 25–35 50–55

10–25 60–70 30–45 20–35 50–70 40–60 65–70 30–60 18–30 50–70 40–55 60–70 50–65 0–5 6–12 35–35 16–25 55–65 40–60 40–50 12–25 22–30 40–47

8.1 Genus Emericella Berk. As noted above, Emericella is an Aspergillus teleomorph characterised by the formation of white cleistothecia surrounded by Hu¨lle cells (thick walled refractile cells like chlamydoconidia) and producing purple ascospores. Conidiophores usually have short brown stipes, bear both metulae and phialides and produce columns of dark green conidia. Christensen and States (1982) accepted 29 species. Christensen and Raper (1978) and Christensen and States (1982) provided keys and descriptions to Emericella species and the related Aspergillus series. Many species are known primarily or solely from desert soils in the western United States and other parts of the world. Only one species is at all common in foods, Emericella nidulans.

Emericella nidulans (Eidam) Vuill.

Fig. 8.2

Anamorph: Aspergillus nidulans (Eidam) G. Winter Colonies on CYA 40–50 mm diam, plane, low, moderately dense to dense, sometimes with a floccose overlay; mycelium white; cleistothecia white, surrounded by white to buff or dull yellow Hu¨lle cells; conidial heads sparse to quite dense, radiate when young, later forming long well defined columns, conidia coloured pale green or when dense dark green;

10–16 8–25 8–12 8–12 25–40 0–10 15–20 18–30 10–16 25–40 20–30 20–40 20–40 8–15 10–15 20–30 15–20 35–40 18–22 10–14 10–18 30–45 20–30

0–25 15–35 10–30 5–20 55–65 65–70 20–50 60–70 30–45 55–65 20–30 40–65 65–70 0 0 0 0–10 50–65 50–70 30–50 0–10 0 0

15–30 68–70 30–45 24–30 65–70 40–60 65–70 60–70 30–50 65–70 45–70 60–70 65–70 0–10 16–20 45–70 25–30 60–70 45–70 25–40 15–30 50–70 45–70

exudate sometimes present, dull red to brown; violet soluble pigment sometimes produced; reverse sometimes pale, usually brightly coloured in shades of orange, orange brown, deep brown or violet brown. Colonies on MEA 35–60 mm diam, occasionally only 25 mm, sometimes low, plane and velutinous with heavy conidial formation and few cleistothecia, sometimes deeper and with abundant cleistothecia surrounded by dull yellow or buff Hu¨lle cells; mycelium white; cleistothecia sometimes abundant, surrounded by Hu¨lle cells, at other times sparsely produced; conidia dark green; reverse pale, brown or violet brown. Colonies on G25N 10–15 mm diam, low and dense; conidia pale green; reverse pale. No growth at 58C. At 378C, colonies 50–70 mm diam, low and sparse, usually predominantly cleistothecial, sometimes with areas of dark green conidia; reverse usually orange or brown. Cleistothecia mostly 100–250 mm diam, white at first but at maturity dark red, maturing on CYA in 8–10 days, surrounded by heavy walled Hu¨lle cells, 15–25 mm diam; ascospores red to purple, ellipsoidal, 4–6 mm long, smooth walled, usually ornamented with two conspicuous longitudinal flanges. Conidiophores borne from aerial hyphae, stipes 60–150 mm long, often sinuous, with smooth, brown walls and with conspicuous footcells; vesicles spathulate to pyriform, 8–12 mm wide, bearing metulae and phialides over the upper half or less, of similar

280

8 Aspergillus and Related Teleomorphs

Fig. 8.2 Emericella nidulans (a) colonies on CYA and MEA, 7 days, 258C; (b, c, d) heads, bars = 10 mm; (e) conidia; and (f) ascospores, bars = 5 mm

size, 6–8 mm long; conidia spherical, green, 3.0–3.5 mm diam, with roughened walls, borne in radiate to loosely columnar heads. Distinctive features. Emericella nidulans is distinguished by growth on CYA at least as fast at 378C as at 258C, developing cleistothecia surrounded by Hu¨lle cells and sparse to abundant green conidia borne on

metulae and phialides from diminutive brown walled stipes. Mature ascospores are red and have either two longitudinal flanges or no ornamentation. Other Emericella species show different ascospore ornamentation. Taxonomy. The nomenclaturally correct name for the anamorph of Emericella nidulans is

8.2

Genus Eurotium

Aspergillus nidulellus Samson and W. Gams (Samson and Gams, 1985). The commonly used name A. nidulans (Eidam) Winter was invalid because it was described inclusive of the teleomorph. However, A. nidulans has remained in common use and is very important to fungal geneticists, so conservation of A. nidulans was sought under the provisions of the Botanical Code (Kozakiewicz et al., 1992). This was rejected by the Special Committee on Fungi and Lichens. Nevertheless, the name A. nidulans was included in the Aspergillus NCU list (Pitt and Samson, 1993) and has been given protected status under the Botanical Code (Greuter et al., 1994). A. nidulans is used here as the appropriate name for the anamorph of E. nidulans. Physiology. According to Panasenko (1967), Emericella nidulans is able to grow from 6–8 to 46–488C, with an optimum at 35–378C. Lacey (1980) gave a range of 10–518C, so this species is a marginal thermophile. E. nidulans germinated at 0.82 aw at 208C (Snow, 1949), 0.80 aw at 378C, 0.81 aw at 258C, 0.83 aw at 208C and 0.90 aw at 158C (Ayerst, 1969), so this species is also a xerophile, a most unusual combination. The heat resistance of E. nidulans ascospores does not appear to have been studied; because of its thermophilic nature, that property might be expected to be high. Mycotoxins. Emericella nidulans has been reported to produce sterigmatocystin, a mycotoxin more commonly associated with Aspergillus versicolor (Frisvad et al., 2006b) and emestrin (Terao et al., 1988, 1990). Emestrin is highly toxic, with an LD50 of 13 mg/kg when given to mice intraperitoneally (Terao et al., 1988). Toxicity due to the growth of E. nidulans has not been reported under practical conditions. Ecology. Although not especially common in foods, Emericella nidulans has been isolated from a wide variety of sources. Cereals and cereal products have been the most common, including wheat, flour and bread, barley, rice and maize and sorghum (see Pitt and Hocking, 1997). Other sources include wine grapes (Bau et al., 2005), peanuts, hazelnuts, meat, soybeans, dried beans, peppercorns, chocolate and spices (see Pitt and Hocking, 1997). We isolated E. nidulans at low levels for Thai soybeans, sorghum and cassava (Pitt et al., 1994). References. Raper and Fennell (1965), as A. nidulans; Christensen and Raper (1978); Domsch et al. (1980); Christensen and States (1982); Klich (2002).

281

8.2 Genus Eurotium Link: Fr. Eurotium is a very well defined and very wellknown genus of Ascomycetes, characterised by the formation of barely macroscopic yellow cleistothecia with smooth, cellular walls. The Aspergillus anamorphs are also characteristic: vesicles bear uncrowded phialides and the phialides produce dull green, spinose conidia in loosely radiate heads. All species of Eurotium are xerophilic and often do not develop characteristic fruiting structures on high water activity media such as CYA and MEA. Incorporation of 20% sucrose into Czapek agar greatly assisted identification of Eurotium species (Thom and Raper, 1941). This medium, of about 0.98 aw, produces suboptimal growth of all Eurotium species but permits development of both anamorphs and teleomorphs plus mycelial colours which greatly aid species identification. Media of lower aw, such as G25N, usually produce rampant growth which is of less value in determinative taxonomy. For compatibility with CYA, we introduced Czapek yeast extract agar with 20% sucrose (CY20S) for Eurotium taxonomy (Pitt and Hocking, 1985). This medium has proved to be of considerable value in general Aspergillus taxonomy (Klich, 2002), but its use here is confined to Eurotium identification. When an isolate has been identified as a Eurotium by using the standard media and incubation conditions, and growth data recorded as usual, the culture should be inoculated onto CY20S agar. After incubation for 7 days at 258C, diameters and colours should be recorded and the cultures returned to the incubator. At intervals, wet mounts of cleistothecia from the central area of a colony should be made, stained with lactofuchsin and examined with the 40 objective. Identification can be completed when mature ascospores are produced, which will be within 14 days for the majority of isolates. Ascospores are mature when they are readily liberated from asci and do not take up stain immediately. Conidia are usually present in wet mounts also and may be distinguished from ascospores by their uniformly spinose walls and the fact that lactofuchsin stains them quite readily. Unstained conidia are green under the microscope while mature ascospores are faintly yellow and are more refractile. Ascospore dimensions should be measured under the 100 objective and

282

8 Aspergillus and Related Teleomorphs

ornamentation noted, i.e. smooth or rough walls and presence or absence of a longitudinal furrow, ridges or flanges. Nomenclaturally correct names were introduced for all common Eurotium anamorphs by Samson and Gams (1985), but they are rarely used. In general, use of the Eurotium names for these species is more logical, as the teleomorphs are often essential for identification and are normally present in Petri dish cultures on CY20S. As described below, differences exist among the anamorphs of Eurotium species, but these are not usually considered in species identification procedures.

About 20 Eurotium species are known (Raper and Fennell, 1965; Blaser, 1975). Descriptions and synonymy for seven species are given by Pitt (1985). Four of those are exceedingly common in all kinds of environments where just sufficient moisture exists to support fungal growth. The three others are encountered less frequently, and the remainder are curiosities known only from three isolates or less. The four common species, E. amstelodami, E. chevalieri, E. repens and E. rubrum, are treated here together with E. herbariorum, which resembles E. rubrum, and E. cristatum, for which E. amstelodami is sometimes mistaken.

Key to Eurotium species included here 1

No growth on CYA, MEA or CY20S; on MY50G cleistothecia white Growth on CY20S; usually growth on CYA and MEA

2 (1)

Ascospores with conspicuous ridges or flanges Ascospores without conspicuous ridges or flanges

3 (2)

Colonies coloured only yellow, from cleistothecia, and green, from conidial heads; ascospores with two prominent, irregular, broad, longitudinal ridges Colonies with conspicuous yellow to orange sterile hyphae; ascospores like pulley wheels, with two prominent, narrow, longitudinal flanges

3 5

4 (3)

Ascospores with rough walls, not exceeding 5 mm long overall Ascospores with walls smooth or very finely spiky, up to 6 mm long overall

5 (2)

Colonies with yellow or orange sterile hyphae; ascospores smooth walled and with just a trace of a longitudinal furrow Colonies with orange to reddish hyphae, in age becoming red brown; ascospores with a fine but distinct longitudinal furrow flanked by two low, minutely roughened ridges

6 (5)

Ascospores up to 6 mm long, asci maturing within 14 days Ascospores frequently exceeding 6 mm long, ascospores usually not maturing within 14 days

Eurotium amstelodami L. Mangin

Fig. 8.3

Aspergillus amstelodami (L. Mangin) Thom & Church (invalid name, includes teleomorph)

Anamorph: Aspergillus vitis Novobr. Colonies on CYA 14–18 mm diam and on MEA 16– 25 mm diam, low and dense, plane; mycelium inconspicuous, white or yellow; abortive yellow cleistothecia conspicuous centrally in patches or sectors,

E. halophilicum (see E. rubrum) 2

4 E. chevalieri E. amstelodami E. cristatum (see E. amstelodami)

E. repens

6 E. rubrum E. herbariorum (see E. rubrum)

surrounded by well formed bright to dark green conidial heads; reverse pale or occasionally dark green. Colonies on G25N 25–35 mm diam, plane, deep and floccose, with appearance usually uniformly dull green from layers of conidial heads; mycelium inconspicuous, white; yellow cleistothecia sometimes visible; conidial heads abundant, dull green; reverse yellow under cleistothecia, or uncoloured. No growth on CYA at 58C. On CYA at 378C, colonies up to 15 mm diam sometimes formed.

8.2

Genus Eurotium

283

Fig. 8.3 Eurotium amstelodami (a) colonies on CYA and CY20S, 7 days, 258C; (b, c) heads, bars = 20 mm; (d) ascospores; and (e) conidia, bars = 5 mm

Colonies on CY20S usually 30–45 mm diam, plane, low and velutinous, usually with a layer of yellow cleistothecia near the agar surface surmounted by a dense layer of radiate dull green conidial heads; reverse uncoloured, i.e. yellow beneath cleistothecia and pale grey green under conidial areas.

Cleistothecia bright yellow, mostly spherical, 75–150 mm diam, usually without surrounding sterile hyphae; maturing in 9–12 days on CY20S and 12–14 days on G25N; ascospores yellow, ellipsoidal, 4.5–5.0 mm long, with rough walls and with two conspicuous, often sinuous, longitudinal ridges of

284

low, irregular height and spacing. Conidiophores borne from aerial hyphae, stipes relatively short, 250–400 mm long, vesicles spherical to spathulate, 18–30 mm diam on CY20S, 35–40 mm on G25N, fertile over the upper two thirds to three quarters, bearing phialides only; phialides 5–8 mm long; conidia spherical to subspheroidal, 4.0–5.0 mm diam, with densely spinulose walls, borne in radiate to loosely columnar heads. Distinctive features. The principal diagnostic feature of Eurotium amstelodami is its conspicuous ascospore ornamentation: wide, irregular ridges or flanges and rough walls. Mycelium remains inconspicuous; colony colours consist only of yellow from the cleistothecia and dull green from the conidia, in both obverse and reverse. Taxonomy. Pitt (1985) pointed out that Mangin’s original concept of this species differed from later usage (Thom and Raper, 1941; Raper and Fennell, 1965), including the neotypification by Blaser (1975). Acceptance of the NCU list for the family Trichocomaceae (Pitt and Samson, 1993) has legitimised the modern concept. The correct anamorph name for this species, Aspergillus vitis, is seldom used (Pitt and Samson, 1993). Physiology. The optimal temperature for growth of Eurotium amstelodami is 33–358C (Domsch et al., 1980), with a maximum at 43– 468C (Blaser, 1975). It was reported to grow down to 0.70 aw at 258C by Armolik and Dickson (1956). Spore germination was noted after 120 days, but not below 0.75 aw within 100 days by Wheeler and Hocking (1988). Scott (1957) showed that the optimal aw for growth was near 0.96 regardless of controlling solute, but the maximum growth rate in sucrose media was twice as fast as in glycerol. Faster growth was observed on media controlled with NaCl than on glycerol based media, and maximum growth rates occurred at 0.90–0.93 aw (Avari and Allsopp, 1983). In a medium of pH 3.8 and 0.98 aw, 80–85% of ascospores of Eurotium amstelodami survived heating at 608C for 10 min; 1–3% survived a similar treatment at 708C; and 0.2% at 758C. Only 0.3% of conidia of Aspergillus vitis survived heating for 10 min at 608C (Pitt and Christian, 1970). Tolerance of sorbate was developed after 21 transfers through increasing concentrations, rising to 0.07% at pH 5.5 (Vin˜as et al., 1990).

8 Aspergillus and Related Teleomorphs

In an atmosphere of 1% O2, Eurotium amstelodami grew at 50% of the rate in air (Hocking, 1990). This species has been reported as one source of chloroanisoles which may cause off odours in food carried in shipping containers (Hill et al., 1995) and of isoprene, a cause of off odour in a bakery product (Berenguer et al., 1991). Ketonic rancidity in coconut has also been caused by E. amstelodami (Kinderlerer, 1984a; Kinderlerer and Kellard, 1984). It showed the highest proteolytic activity on meat of any Eurotium species (Binzel, 1980) and was relatively resistant to propionic acid (Mu¨ller et al., 1981), but lipase activity was low (Magan et al., 1993). Marı´ n et al. (2003) reported N-acetyl-b-Dglucosaminidase activity at 0.90 aw at pH 6 and at 0.85 aw at pH 7.5. Conidia of E. amstelodami were found to survive in 30% (w/v) NaCl solution over a period of 14 weeks (Butinar et al., 2005b). Mycotoxins. Isolates of Eurotium amstelodami from hams were toxic to chick embryos and when injected into mice (Resurreccion and Koehler, 1977). However, the possible toxins involved have not been identified (Frisvad and Samson, 1991), and it is generally assumed that this is a benign fungus, like most Eurotium species. Ecology. In Australia, E. amstelodami is less common than the other Eurotium species described here; however, literature reports indicate that elsewhere it is a ubiquitous foodborne species, usually associated with stored goods. This species has been reported to spoil rapeseed due to heat damage, Japanese traditional noodles and cheese (see Pitt and Hocking, 1997). E. amstelodami has been reported as one cause of gushing in beer (Gyllang and Martinson, 1976). As with other Eurotium species, cereals are a major substrate, including wheat, flour, refrigerated dough, bread and bakery products, rice and rice bran, barley, maize, cereal flakes and corn snacks (see Pitt and Hocking, 1997). Other sources include grapes and raisins (Valero et al., 2007a), mixed feeds, cereals and legume feeds (Accensi et al., 2004), pasta (Halt et al., 2004), peanuts, hazelnuts and walnuts, pistachios, soybeans, sunflower seeds, meat products, smoked bacon, biltong, dried salt fish and jam (see Pitt and Hocking, 1997). Eurotium amstelodami was less common in Southeast Asian commodities than E. chevalieri or E. rubrum. Nevertheless, we isolated it at 1% or more of all particles examined in maize, peanuts, soybeans,

8.2

Genus Eurotium

285

cashews, copra, paddy rice, mung beans, sorghum and peppercorns. Percentages of samples infected were often relatively low, usually 8–15%, but rates of infection in infected samples were often high, up to 40–60%. This indicates some samples becoming infected during storage (Pitt et al., 1993, 1994, 1998a). Additional species. Eurotium cristatum (Raper and Fennell) Malloch and Cain, anamorph Aspergillus cristatellus Kozak., synonym E. chevalieri var. intermedius Thom and Raper (invalid name): in culture this species resembles E. amstelodami and is often mistaken for it. However, ascospores are larger, up to 6 mm long including crests, and have walls ornamented with discrete, very fine spikes. Unlike E. amstelodami, colonies have bright yellow hyphae surrounding the cleistothecia. No physiological studies are known, and mycotoxin production is also unknown. We have isolated this species uncommonly, from heated strawberry puree and Philippine peanuts. References. Eurotium amstelodami: Raper and Fennell (1965), as Aspergillus amstelodami; Blaser (1975); Domsch et al. (1980); Pitt (1985); Klich (2002). E. cristatum: Blaser (1975); Pitt (1985).

Eurotium chevalieri L. Mangin

Fig. 8.4

Anamorph: Aspergillus chevalieri L. Mangin Colonies on CYA 16–25 mm diam, low and dense, plane or lightly sulcate; mycelium bright yellow, often darker centrally, enveloping abundant abortive yellow cleistothecia and overlaid by sparse to abundant greyish green conidial heads; yellow brown soluble pigment sometimes produced; reverse pale to orange or deep brown. Colonies on MEA 16–25 mm diam, plane, dense to floccose; mycelium white to yellow, overall colours and characteristics as on CYA; yellow soluble pigment sometimes produced; reverse pale, olive, orange or brown. Colonies on G25N 20–30 mm diam, usually very deep and floccose, with pale to bright yellow mycelium enveloping abundant developing cleistothecia and overlaid by sparse to abundant greyish green conidial heads, occasionally development more sparse and producing few cleistothecia; reverse usually pale, sometimes yellow or dull green. No growth on CYA at 58C. Usually no growth on CYA at 378C, occasionally colonies up to 10 mm diam formed.

Colonies on CY20S 45–65 mm diam, plane, low and dense; colony character varying with production of conidial heads, in isolates with sparse heads, mycelium white at the margins, then bright to deep yellow or orange centrally, enveloping abundant cleistothecia, in isolates with heavy conidial production these elements more or less obscured and colony appearance dominated by conidia, dull green; reverse pale, yellow, orange brown or dull green. Cleistothecia bright yellow, spherical, 100–140 mm diam, enveloped in yellow to orange vegetative hyphae, maturing at colony centres in 8–10 days on CY20S and 12–14 days on G25N; ascospores yellow, ellipsoidal, shaped like pulley wheels, 4.5–5.0 mm long, smooth walled, with two prominent, parallel, narrow, sometimes sinuous, longitudinal flanges. Conidiophores borne from aerial hyphae, stipes mostly 400–700 mm long, thin walled, already collapsing in 7 days, broadening to vesicles 25–35 mm diam, fertile over the whole area, bearing phialides only; phialides ampulliform, 5–8 mm long; conidia on CY20S and G25N ellipsoidal or doliiform (barrel shaped), 4.0–5.5 mm long, with spinose walls. Distinctive features. As with other Eurotium species, the ascospore is the prime distinguishing feature: in E. chevalieri these are characteristically shaped like minute pulley wheels. Colonies grow quite well on CYA and MEA, but sporulate poorly; colonies on CY20S produce conspicuous yellow to orange hyphae, while on CYA and MEA they can be brown. Taxonomy. This taxon was originally described as a holomorph species, so the name ‘‘chevalieri’’ is valid for both the Eurotium and the Aspergillus morphs (Pitt and Samson, 1993). Physiology. The optimum temperature for growth of Eurotium chevalieri is 30–358C (Domsch et al., 1980), with a maximum at 40–438C (Blaser, 1975). Ayerst (1969) obtained growth down to 0.71 aw at 338C; Pitt and Christian (1968) reported a minimum of 0.74 aw at 258C on a medium of pH 3.8. Germination of ascospores was little affected by glucose or glycerol as solute, but was slower in NaCl. Media of pH 4 or 6.5 did not affect germination. Maximum growth rates were much higher on glucose/fructose or NaCl media than in the presence of glycerol; again pH had little effect. The optimum aw for growth was 0.94–0.95 (Pitt and Hocking, 1977). E. chevalieri was able to grow in the presence of 2,000 ppm potassium sorbate, following a 20 day

286

8 Aspergillus and Related Teleomorphs

Fig. 8.4 Eurotium chevalieri (a) colonies on CYA and CY20S, 7 days, 258C; (b, c) heads, bars = 10 mm; (d) ascospores; and (e) conidia, bars = 5 mm

8.2

Genus Eurotium

lag period, at 0.85 aw and pH 6.0. No growth was recorded at lower pH and water activity values (e.g. 0.80 aw and pH 5.5) in the presence of preservative (Char et al., 2006). Eurotium chevalieri was the most heat resistant xerophilic fungus studied by Pitt and Christian (1970): 18–25% of ascospores survived heating at 708C for 10 min in a medium of 0.98 aw and pH 3.8, and up to 0.5% a similar treatment at 808C. The decimal reduction line had an F80 of 3.3 min with a high z value, 12.8C8, under these conditions. However, conidia of Aspergillus equitis were not especially heat resistant: only 0.1% survived heating for 10 min at 608C. This species has been reported as one source of isoprene, a cause of off-odour in a bakery product (Berenguer et al., 1991). It is also a cause of ketonic rancidity in coconut (Kinderlerer, 1984a; Kinderlerer and Kellard, 1984). High proteolytic activity on meat (Binzel, 1980) and moderate lipolytic activity on vegetable oils (Kuku, 1980) have been reported. Mycotoxins. Eurotium chevalieri has been reported to produce toxic compounds, identified as echinulin and neoechinulin, which cause feed refusal in swine (Vesonder et al., 1988). However, other tests for toxicity of E. chevalieri have been negative (Frisvad and Samson, 1991; Adebajo and Oyesiku, 1994). Ecology. Eurotium chevalieri must rank as one of the most common spoilage fungi on earth, especially in warmer regions. It has been reported to cause spoilage of high moisture prunes, pecans, Japanese traditional noodles, a semisoft baked cookie and, in our laboratory, cheese spread, chick peas and faba beans (see Pitt and Hocking, 1997). Like the other common species in this genus, it has been isolated from a great variety of foods, especially cereals, including wheat and flour, rice, rice flour and rice bran, maize and cereal flakes. It is also common in processed and dried meats and nuts: peanuts, pecan nuts, hazelnuts and walnuts and coconut. Other sources include filled chocolates, dried beans and peas, soybeans, sunflower seeds, cocoa beans, spices, dried salt and cured fish (see Pitt and Hocking, 1997). It has recently been isolated from evaporated milk products in Argentina (Char et al., 2005) and black pepper in Brazil (Gatti et al., 2003). Many other habitats no doubt exist. This species was commonly isolated in our study of Southeast Asian commodities, particularly from peanuts and maize from Thailand, Indonesia and the

287

Philippines. E. chevalieri infections in other nuts and oilseeds were also heavy (Pitt et al. 1993, 1994, 1998a). Infections in beans were usually much lower, except for soybean samples from the Philippines (Pitt et al., 1993, 1994, 1998a). Rice was less affected: no more than 20% of samples were infected, and overall infection rates in milled rice were only 2–4% (Pitt et al., 1993, 1994, 1998a). References. Raper and Fennell (1965), as Aspergillus chevalieri; Blaser (1975); Domsch et al. (1980); Pitt (1985); Klich (2002).

Eurotium repens de Bary

Fig. 8.5

Aspergillus repens (Corda) de Bary Eurotium pseudoglaucum (Blochwitz) Malloch & Cain Aspergillus pseudoglaucus Blochwitz

Anamorph: Aspergillus reptans Samson & W. Gams Colonies on CYA and MEA 10–20 mm diam, plane, deep and dense, mycelium white, yellow or orange, either enmeshing abortive yellow cleistothecia or surmounted by dull green to dull blue conidial heads, or both, depending on isolate; reverse pale, dull yellow, green or orange, less commonly bright yellow or orange. Colonies on G25N usually 25–45 mm diam, plane, deep and floccose, sometimes reaching the Petri dish lid; mycelium white to bright yellow or orange, usually enmeshing many layers of developing bright yellow cleistothecia and overlaid by sparse dull green conidial heads, in uncommon isolates cleistothecia and yellow hyphae inconspicuous and dull green conidial heads dominant; reverse pale, brilliant yellow to orange, or orange brown. No growth on CYA at 5 or 378C. Colonies on CY20S 45–65 mm diam, plane, low or somewhat floccose, broader but much less luxuriant than on G25N; mycelium white to yellow or orange, overall colour varying from yellow with scattered dull green areas in predominantly cleistothecial isolates to dull green or bluish green in those with conidial heads dominant; reverse dull green or bright yellow to orange or both, at maturity dull yellow brown. Cleistothecia on CY20S or G25N borne from and enveloped in sterile yellow to orange hyphae, bright yellow, spherical, 75–100(–125) mm diam, maturing at colony centres in 7–10 days; ascospores yellow, ellipsoidal, 5.0–5.5 mm long, without ridges

288

8 Aspergillus and Related Teleomorphs

Fig. 8.5 Eurotium repens (a) colonies on CYA and CY20S, 7 days, 258C; (b, c) heads, bars = 10 mm; (d) ascospores; and (e) conidia, bars = 5 mm

or flanges and with no more than a trace of a longitudinal furrow, smooth walled, borne in loosely radiate heads. Conidiophores borne from aerial hyphae, stipes mostly 500–1,000 mm long,

broadening to vesicles 15–30 mm diam, fertile over the whole area, bearing phialides only; phialides ampulliform, 7–10 mm long; conidia on CY20S spherical to subspheroidal, 5–7 mm diam, on

8.2

Genus Eurotium

G25N ellipsoidal to pyriform, 7–10 mm long, with densely spinulose to spinose walls. Distinctive features. The prime distinction of Eurotium repens from other Eurotium species is the production of ascospores without ridges or flanges, usually without a longitudinal furrow, and smooth walls. Hyphal and reverse colours on CYA and MEA are yellow to orange, never red. Taxonomy. Samson and Gams (1985) provided a correct anamorph name for this species, Aspergillus reptans, but it is rarely used. Physiology. Eurotium repens grows between 4–58C and 38–408C, with an optimum at 25–278C (Panasenko, 1967; Gonzalez et al., 1988). The minimum aw for germination of E. repens was reported as 0.72 on media of neutral pH, at temperatures of 20– 258C (Snow, 1949; Armolik and Dickson, 1956; Magan and Lacey, 1984a). Andrews and Pitt (1987) reported that the minimum aw for germination in NaCl and glycerol media were 0.83 and 0.72 aw, with optima at 0.95 and 0.91 aw, respectively. In media controlled by glucose/fructose, the minimum aw for germination was much lower, 0.69 aw (Andrews and Pitt, 1987). The influence of pH from 4.0–6.5 was slight (Avari and Allsopp, 1983), with pH 4.5–5.5 being optimal (Gock et al., 2003). Eurotium repens ascospores are tolerant of elevated temperatures and survived heating to 60 and 708C for 10 min; however, they did not recover after heating at 758C for 10 min (Pitt and Christian, 1970). Eurotium repens has been reported as one source of chloroanisoles which may cause off-odours in food carried in shipping containers (Hill et al., 1995). It is one of a group of species producing odorous compounds including 3-methyl-1-butanol, 1-octen-3-ol and 3-octanone in grain, useful in quality monitoring systems (Sinha et al., 1985). Mycotoxins. Mycotoxins are not known to be produced by this species (Frisvad and Samson, 1991). Ecology. Eurotium repens is a very common species. It has been reported to cause spoilage of cheese, cheesecake, pecans, maize and traditional Japanese noodles. It is very common in stored grains, for example, wheat, barley, maize and rice. E. repens is of common occurrence on processed and dried meat or fish, for example salami, meat products, ripening and ripened hams, biltong, dried salted fish and katsuobushi. Other sources include nuts, sunflower seed and red pepper powder (see Pitt and Hocking, 1997).

289

In our laboratory, Eurotium repens has been isolated from spoiled prunes, salami, strawberry puree, bread, cake, nuts, cheese and other products. It was relatively uncommon in Southeast Asian commodities by comparison with E. chevalieri or E. amstelodami, exceeding 1% infection overall only in copra from Thailand (3%) and kemiri nuts from Indonesia (2%) (Pitt et al., 1993, 1994, 1998a). Eurotium repens has found use in food manufacture, as a starter culture in the manufacture of katsuobushi from bonito (Dimici and Wada, 1994) and of fish sauce from fish meal (Hayakawa et al., 1993). References. Raper and Fennell (1965) as Aspergillus repens; Blaser (1975); Domsch et al. (1980); Pitt (1985); Klich (2002).

¨ Eurotium rubrum Jos. Konig et al.

Fig. 8.6

Aspergillus sejunctus Bainier & Sartory (invalid name, includes teleomorph) Aspergillus ruber (Jos. Konig et al.) Thom & Church (invalid ¨ name, includes teleomorph)

Anamorph: Aspergillus rubrobrunneus Samson & W. Gams Colonies on CYA 10–20 mm diam, plane, deep, usually dense and velutinous, sometimes floccose; mycelium yellow to bright orange; conidiophores and developing cleistothecia usually present, but often poorly formed; reverse pale yellow to orange brown. Colonies on MEA usually 10–20 mm diam, sometimes only 5–8 mm, similar to those on CYA, but sometimes less deep and with more conspicuous orange hyphae. Colonies on G25N 30–45 mm diam, plane, often floccose and with hyphal strands sometimes reaching the Petri dish lid, consisting of cleistothecia in layers supported and surrounded by relatively sparse orange to dark orange hyphae; conidial heads usually rare, above or within the cleistothecial layer, dull green; reverse yellow, brown or reddish orange. No growth on CYA at 5 or 378C. Colonies on CY20S at 7 days 35–60 mm diam, plane or lightly sulcate, usually low, dense and velutinous; mycelium conspicuous, at the margins yellow, becoming orange or more reddish elsewhere, enveloping abundant yellow cleistothecia and surmounted by sparse to abundant dull green heads; reverse in colours similar to the mycelium, though rarely so bright, or deep yellow brown; in

290

8 Aspergillus and Related Teleomorphs

Fig. 8.6 Eurotium rubrum (a) colonies on CYA and CY20S, 7 days, 258C; (b, c) heads, bars = 10 mm; (d) mature cleistothecium releasing ascospores, bar = 25 mm; and (e) conidia (spiny) and ascospores, bar = 5 mm

age, 10 days or more, developing bright to brick red brown hyphal colours over the whole plate or in patches, not under densely conidial areas or, rarely, remaining orange; reverse darkening over time to deep brown or almost black. Cleistothecia on CY20S or G25N borne from and enveloped in sterile orange to red hyphae, spherical, yellow, 80–150 mm diam, maturing in 9–12 days; ascospores ellipsoidal, yellow, 5.0–6.0 mm long, with a shallow longitudinal furrow flanked by low, usually minutely roughened ridges, otherwise with

smooth walls. Conidiophores borne from aerial hyphae, stipes mostly 300–700 mm long, terminating in spherical vesicles, 20–35 mm wide, fertile over the upper two-thirds, bearing phialides only; phialides 7–9 mm long; conidia subspheroidal to ellipsoidal, less commonly spherical or pyriform, generally 6–7 mm long, larger in occasional isolates, with spinose walls, borne in loosely radiate heads. Distinctive features. Colonies of Eurotium rubrum on CY20S usually show areas of brilliant red or rusty colours after 10 days or more incubation.

8.2

Genus Eurotium

Ascospores are 5.0–6.0 mm long, rarely otherwise, and show a definite longitudinal furrow and low, minutely roughened ridges. Taxonomy. The commonly used anamorph name Aspergillus ruber and A. sejunctus, which predates it, are both invalid because they include the teleomorph. Samson and Gams (1985) provided a correct anamorph name, A. rubrobrunneus, but it is rarely used. Physiology. Growth temperatures for Eurotium rubrum are probably similar to those reported for E. repens: minimum ca. 58C, optimum 25–278C and maximum near 408C (Panasenko, 1967). The minimum aw for germination has been reported as 0.70 aw (Snow, 1949; Gock et al., 2003) and 0.72 aw (Armolik and Dickson, 1956), at 258C, or 0.72 at 208C and 0.73 at 258C (Wheeler et al., 1988b). On a medium of pH 3.8, germination occurred down to 0.75 aw after 98 days at 258C (Pitt and Christian, 1968). Maximum growth was achieved at 308C at 0.92 aw (Gock et al., 2003). E. rubrum was able to grow on media with up to 20% (w/v) NaCl (0.87 aw) and germination occurred at 25% (w/v) NaCl (0.82 aw), although mycelial growth did not follow (Butinar et al., 2005b). Growth rates were similar in glycerol at pH 4.0 and 6.5, and NaCl at pH 6.5, over a wide aw range; growth in NaCl at pH 4.0 was much slower (Wheeler et al., 1988b). The optimum aw for growth was ca. 0.94 and varied little over all conditions tested (Avari and Allsopp, 1983). At higher temperatures on glucose/fructose media of pH 6.5, the minimum aw was 0.76 at 308C and 0.79 at 378C (Wheeler et al., 1988b). On NaCl-based media, the minimum aw for germination was 0.86–0.88 aw from 20–348C; at 378C it was 0.91 aw, indicating temperature had little effect (Wheeler et al., 1988b). E. rubrum was a highly competitive species over a wide range of aw and temperature on glucose based media, but not in NaCl based media (Wheeler and Hocking, 1993). Eurotium rubrum was relatively tolerant of modified atmospheres, producing visible colonies in 65% CO2 + 1% O2 at 0.90 aw in 15 days. At lower aw values, growth was inhibited: at 0.82 aw and 50% CO2 + 1% O2, visible growth occurred after 35 days, and at 0.78 aw, growth occurred in 20% CO2 + 1% O2 after 25 days. Growth did not occur in higher concentrations of CO2 at these aw values (our unpublished data). A high percentage (80–100%) of ascospores of Eurotium rubrum survived heating at 608C for

291

10 min, at aw 0.98 and pH 3.8; 0.5% or less survived 10 min at 708C and there were no survivors after 10 min at 758C. Conidia of Aspergillus rubrobrunneus were much less heat resistant: only 8% survived 10 min at 508C, 3% 10 min at 608C and none at 708C under the same conditions (Pitt and Christian, 1970). This species developed tolerance to sorbate after 21 transfers through increasing concentrations, rising to 0.07% at pH 5.5 (Vin˜as et al., 1990). Mycotoxins. Several reports have been published which indicate that Eurotium rubrum produces a range of toxic compounds. However, well documented confirmation of toxicity is still lacking (Frisvad and Samson, 1991). Production of echinulin and flavoglaucin by E. rubrum was not correlated with pH or water activity. Further, metabolite production was enhanced on media containing 0.3% propionate (Suhr and Nielsen, 2004). Ecology. A very widely distributed xerophilic fungus, Eurotium rubrum, appears to be somewhat more common in warmer regions. It has caused spoilage of dried and high moisture prunes, coconut and pancake syrup. It has been reported from a wide range of cereals (Lugauskas et al., 2006), especially wheat, maize and rice. It is also common on nuts, meat products and dried salt fish. Other reports have been from sunflower seeds, sesame seeds, cocoa beans, palm kernels, dried vegetables and jam (see Pitt and Hocking, 1997). It has frequently spoiled fruit cakes in Australia (our observations). E. rubrum has been isolated from salterns in the Adriatic (Butinar et al., 2005b). Along with Eurotium repens, E. rubrum has found use in food manufacture, as a starter culture in the manufacture of katsuobushi from bonito (Dimici and Wada, 1994). Eurotium rubrum was readily isolated from most food commodities we examined from Southeast Asia. It was particularly common in peanuts, kemiri nuts, cow peas and red kidney beans, mung beans, soybeans and maize kernels, with high levels in some soybeans. Levels in milled rice were low, however (Pitt et al., 1993, 1994, 1998a). Additional species. Eurotium herbariorum Link [anamorph Aspergillus glaucus Link, synonym A. manginii Raper and Thom (invalid name)] resembles E. rubrum in many features. It is distinguished by the following: (i) colonies on CYA and MEA at 258C do not exceed 10 mm diam in 7 days and are sometimes absent; (ii) colonies on CY20S and

292

G25N rarely exceed 35 mm diam in 7 days; (iii) cleistothecia on CY20S and G25N develop slowly, with ascospores usually taking more than 14 days to mature; and (iv) while ascospores are similar in appearance to those of E. rubrum, they are larger, commonly 6–7(–8) mm long. Taxonomic confusion exists because some authors have synonymised Eurotium rubrum with E. herbariorum or E. repens with E. herbariorum. In our opinion these three species are distinct, although closely related. Eurotium herbariorum is a vigorous xerophile. When grown on media of pH 3.8 containing glucose/fructose as the controlling solute, ascospores of E. herbariorum germinated at 0.74 aw after 19 days, the shortest lag time of any of the common species. Conidia of Aspergillus glaucus germinated at 0.75 aw in 14 days (Pitt and Christian, 1968). When heated in 58Brix grape juice, E. herbariorum ascospores showed a D70 of 2.5 min and a z value of 9.1C8; in 658Brix concentrate, the D70 was 5.2 min and z value 7.1C8 (Splittstoesser et al., 1989). Eurotium herbariorum is able to grow in low oxygen concentrations, producing visible colonies at 258C in 0.5% O2 at 0.82 aw within 60 days and at 0.78 aw within 75 days. It also tolerated elevated CO2 concentrations at reduced aw: at 0.82 aw producing visible colonies in 20 days in 35% CO2 + 1% O2 and 34 days in 50% CO2 + 1% O2. At 0.78 aw, visible growth occurred in 35% CO2 + 1% O2 only after 95 days, and no growth occurred in 50% CO2 + 1% O2 (our unpublished data). Although a far less common species than Eurotium rubrum, E. herbariorum is nevertheless widespread. Spoilage by E. herbariorum has occurred in French and Australian prunes and European cheese (see Pitt and Hocking, 1997). It has also been recorded from maize, meat products, rice, sardines and miso and spices (see Pitt and Hocking, 1997). We isolated it from samples of sorghum and peanuts from Indonesia and from one of velvet beans (30% infection). Low levels were present in coriander from Indonesia and soybeans and mung beans from Thailand (Pitt et al., 1993, 1994, 1998a). Additional species. Eurotium halophilicum C.M. Chr. et al. is distinguished by the production of white cleistothecia and by inability to grow on CYA, MEA or CY20S, even after 14 days incubation. On MY50G, after 14 days, colonies are 15–20 mm diam, consisting almost entirely of cleistothecia

8 Aspergillus and Related Teleomorphs

surrounded by a weft of white mycelium (Hocking and Pitt, 1988). Aspergillus heads, characteristic of those from Eurotium species, are formed only on concentrated media, below 0.85 aw, and bear pyriform, rough walled conidia 8–11 mm long. It was originally described as a halophile (Christensen et al., 1959), but in fact is an extreme xerophile, failing to germinate above 0.94 aw, with growth occurring optimally between 0.85 and 0.80 aw. The minimum recorded germination occurred at 0.68 aw after only 38 days incubation at 258C (Andrews and Pitt, 1987). Germination occurred on saturated NaCl based media (0.75 aw) in a similar time. This is undoubtedly a rare species, but is of interest because it will not be isolated on any medium designed for the recovery of Eurotium species. It is also very difficult to recognise because its cleistothecia are white and it produces its anamorph only at very low aw. In our laboratory we isolated it from longstored cardamom seeds on MY50G, the medium usually reserved for Xeromyces bisporus and xerophilic Chrysosporium species (Hocking and Pitt, 1988). References. Raper and Fennell (1965), Eurotium rubrum under Aspergillus ruber and E. herbariorum under A. manginii; Pitt (1985), E. rubrum and E. herbariorum; E. herbariorum, Klich (2002); E. halophilicum, Hocking and Pitt (1988).

8.3 Genus Neosartorya Malloch and Cain Neosartorya produces cleistothecia with cellular walls like Eurotium; however, the walls and ascospores are colourless or white, not yellow. Like those of Eurotium, Neosartorya anamorphs produce vesicles bearing phialides only, but vesicles are small and pyriform, enlarging towards the apices. Twelve species and varieties are currently accepted (Kirk et al., 2001), which mainly inhabit soil and decaying vegetation. All are thermoduric or thermophilic, none are xerophiles. The well known human pathogen Aspergillus fumigatus is closely related (Girardin et al., 1995; Geiser et al., 1998), and occasional pathogenicity to humans has been reported. Neosartorya isolates should be handled with care. From the viewpoint of the food technologist, the main importance of Neosartorya species is the very high heat resistance of their ascospores. Neosartorya fischeri, described below, is the only Neosartorya species significant in foods.

8.3

Genus Neosartorya

Neosatorya fischeri (Wehmer) Malloch & Cain

293

Fig. 8.7

Aspergillus fischeri Wehmer (invalid name, includes teleomorph)

Anamorph: Aspergillus fischerianus Samson & W. Gams

Colonies on CYA 50–60 mm diam, occasionally larger, plane, sparse to moderately dense, surface texture floccose; mycelium white to pale yellow, enveloping abundant developing cleistothecia; conidial heads usually small and sparsely produced, above the layer of cleistothecia, grey green; reverse pale to

Fig. 8.7 Neosartorya fischeri (a) colonies on CYA and MEA, 7 days, 258C; (b, c) heads, bars = 10 mm; (d) ascospores; and (e) conidia, bars = 5 mm

294

pinkish brown, sometimes yellow centrally. Colonies on MEA 60 mm or more diam, usually covering the whole Petri dish, low and sparse; mycelium white to pale yellow, surrounding layers of developing white cleistothecia; conidial heads small and sparse, grey green; reverse pale, pale brown or dull yellow. Colonies on G25N 8– 12 mm diam, of sparse to dense white mycelium; reverse pale to dull yellow. No growth at 58C. At 378C, colonies covering the whole Petri dish, plane or sulcate, of white or grey mycelium and abundant conspicuous white cleistothecia; reverse pale to pinkish brown. Cleistothecia 150–400 mm diam, with a definite wall of flattened cells with hyphae attached, white to cream, maturing in 9–12 days at 258C, slightly faster at 378C; ascospores ellipsoidal, 7–8 mm long overall, including two prominent, sinuous longitudinal flanges, other irregular ridges sometimes present as well. Conidiophores borne from aerial mycelium, stipes usually 300–500 mm long, with thin, colourless, smooth walls, enlarging terminally to pyriform vesicles; vesicles 12–18 mm diam, fertile over the upper half, bearing phialides only; phialides 5–6 mm long, with short necks; conidia subspheroidal to ellipsoidal, 2.5–3.0 mm long, smooth walled, borne in radiate to columnar heads. Distinctive features. Colonies of Neosartorya fischeri spread rapidly at both 25 and 378C and are white; white cleistothecia and inconspicuous grey green Aspergillus heads are produced. Taxonomy. Earlier literature discussed Neosartorya fischeri under its invalid anamorph name Aspergillus fischeri. However, the teleomorph is produced under most growth conditions and, from the food technologist’s point of view, is the morph responsible for this species’ most important property, high heat resistance. N. fischeri is thus the appropriate name for this species when it is found in foods. Samson and Gams (1985) provided a valid anamorph name, A. fischerianus. Two varieties of Neosartorya fischeri were accepted by Raper and Fennell (1965), distinguished by differences in ascospore ornamentation. These varieties were raised to species status by Kozakiewicz (1989), as N. glabra (Fennell and Raper) Kozakiewicz and N. spinosa (Fennell and Raper) Kozakiewicz, again based on the

8 Aspergillus and Related Teleomorphs

differences in ascospore ornamentation. Other differences between these taxa appear to be very small. So, in agreement with Samson et al. (1990), we have not followed Kozakiewicz (1989) here, but maintained N. fischeri as a species with well defined varieties. It is worth noting that only N. fischeri var. glabra and N. fischeri var. spinosa normally are found in foods. That fact suggests ecological differences between these taxa, and if that is confirmed, may provide a basis for maintaining separation at species level. Molecular studies have shown a very close relationship between N. fischeri var. glabra and N. fischeri var. spinosa and between N. fischeri var. fischeri and A. fumigatus (Girardin et al., 1995). Physiology. Ascospores of this species rank with those of Byssochlamys as the most heat resistant fungal spores known. Kavanagh et al. (1963) reported that ascospores of an isolate more recently identified as Neosartorya fischeri withstood boiling in distilled water for 60 min. They reported that spore age, pH and sugar concentration affected heat resistance. McEvoy and Stuart (1970) heated ascospores of N. fischeri in distilled water: they reported 100% survival after 20 min at 808C and 0.002% survival after 5 min at 1008C. This degree of heat resistance is comparable with that of many bacterial spores and is higher than that of Byssochlamys fulva ascospores. More detailed studies of the heat resistance of N. fischeri ascospores indicate D88 values of 1.2–7.5 min (Quintavalla and Spotti, 1993) or of 4.2–16.2 min (Beuchat, 1986) or a D88 value of 1.4 min and a z value of 5.6C8 (Scott and Bernard, 1987). A mathematical model for the combined effect of aw, pH and redox potential on the heat resistance of N. fischeri has been published (Reichart and Mohacsi-Farkas, 1994). Saloma˜o et al. (2007) reported that citric acid had a greater influence than malic acid on the thermal resistance of ascospores of N. fisheri. Ninety day old ascospores, isolated from tomatoes, were successfully killed by a heat treatment of 1158C for 30 s (Pacheco and De Massaguer, 2004). High pressure treatments (600–900 MPa) for 20 min at 208C in apricot nectar reduced counts of Neosartorya fischeri 100 fold. At 508C, inactivation required less than 4 min at 800 MPa and at 608C, 1–2 min at 700 MPa. Pressure resistance in distilled

8.4

Genus Aspergillus

water was lower (Maggi et al., 1994). Pressure resistance of ascospores increases with age: ascospores that were 3 weeks old were relatively sensitive to pressure treatment (600 MPa for 10 min), with pressure resistance increasing as ascospores matured, at least up to 15 weeks (the oldest ascospores tested). Ascospores of N. fischeri var. spinosa that were 9–15 weeks old were activated by a pressure treatment of 600 MPa for 10 min (Chapman et al., 2007). Outbreaks of spoilage due to this species seem to be rare. In view of its exceptional heat resistance, we speculated that this may be due to an inability to grow at low oxygen tensions. However, growth has been reported in O2 levels as low as 0.1% at 258C (Nielsen et al., 1989). Mycotoxins. The production of fumitremorgens A and C and verruculogen by this species has been confirmed (Beuchat et al., 1988; Nielsen et al., 1988; Frisvad and Samson, 1991). However, although these compounds are highly toxic, this is unlikely to have practical significance, as Neosartorya fischeri has only rarely been reported as growing in foods. Ecology. Neosartorya fischeri ascospores have been isolated from canned strawberries, which had been processed at 1008C for 12 min (Kavanagh et al., 1963). Isolation of N. fischeri from pasteurised fruit juices and fruit powders has been reported several times, but only occasionally from spoiled product (see Pitt and Hocking, 1997). On several occasions we have isolated Neosartorya fischeri from pasteurised fruit juices, strawberry purees and syrups, and a sports drink, but have only seen actual spoilage in canned strawberry puree. Heat treated fresh fruit frequently yield isolates of this species (Spotti et al., 1992, and our observations). Neosartorya fischeri has been reported only rarely from foods which have not been heat treated or processed: from pistachios, soybeans and fresh mangoes (see Pitt and Hocking, 1997). Contamination from soil rather than fungal growth appears probable. N. fischeri has been reported from a case of pulmonary aspergillosis in a liver transplant patient (Gori et al., 1998). References. Raper and Fennell (1965); Domsch et al. (1980); Hocking and Pitt (1984); Klich (2002).

295

8.4 Genus Aspergillus Fr.: Fr. Aspergillus is among the best known and most frequently recognised fungal genera on earth. Species of Aspergillus are among the most economically important fungi, on the positive side being very widely used for synthesis of chemicals, for biosynthetic transformations and enzyme production. On the negative side, they are of great importance in food spoilage and they produce the aflatoxins, the most important of all mycotoxins. Current taxonomies recognise about 185 Aspergillus species (Kirk et al., 2001). Perhaps 30 of these are well defined and usually readily distinguished. Peripheral to these central, common species are a large number of variations on each theme, often with the status of species, but which are seldom encountered and are little more than ephemeral variants. Placement of an unknown isolate in the central species is sufficient identification in all but the most detailed investigations. Twenty four species are described below. Together with the 8 teleomorph species described earlier in this chapter, these 32 represent the vast majority of Aspergilli which will be isolated from foods. These species are all keyed out in the general key at the beginning of this chapter, and the Aspergillus species are described in alphabetical order.

Aspergillus aculeatus lizuka

Fig. 8.8

Aspergillus japonicus var. aculeatus (Iizuka) Al-Musallam

Colonies on CYA 65–70 mm diam or more, plane, velutinous, sometimes with a central floccose overlay; mycelium white; conidial heads red brown to black; pinkish sclerotia sometimes present; exudate and soluble pigment absent; reverse drab yellow to brown. Colonies on MEA 45–70 mm diam, plane, velutinous, mycelium white; conidial heads rather sparse, reddish brown or black; reverse uncoloured or yellowish. Colonies on G25N 15–20 mm diam, plane, sparse, velutinous, similar to those on CYA; reverse pale. No growth at 58C. At 378C, colonies 5– 30 mm diam, consistently less than half of the diameter on CYA at 258C. Conidiophores borne from subsurface or surface hyphae, stipes 500–2,000 mm long, smooth walled, sometimes brown; vesicles 25–60 mm diam, bearing phialides only, covering three quarters of the vesicle

296

8 Aspergillus and Related Teleomorphs

Fig. 8.8 Aspergillus aculeatus Iizuka: (a) colonies on CYA and MEA, 7 days, 258C; (b) heads, bar = 20 mm; (c) head showing the presence of phialides only, bar = 10 mm; (d) conidia, bar = 5 mm

8.4

Genus Aspergillus

surface or more, 5–9 mm long; conidia ellipsoidal, sometimes subspheroidal, 4–5 mm long, spinose, borne in radiate heads, in age splitting into columns. Distinctive features. Red brown to black colonies indicate a close relationship with Aspergillus niger; however, vesicles of A. aculeatus bear phialides only, unlike those of A. niger. A. japonicus also produces phialides only, but A. aculeatus is distinguished from it by the production of larger vesicles and ellipsoidal conidia. Taxonomy. Aspergillus aculeatus and A. japonicus are very closely related species, and A. aculeatus has sometimes been considered to be a variety of A. japonicus (Al-Musallam, 1980). These two species have morphological differences (Raper and Fennell, 1965) and differences in secondary metabolites and RFLP profiles that correlate with each other (Parˇ enicova´ et al., 2001). However, a study to correlate the morphological differences with these other parameters has not been carried out. ITS sequences of the two species are essentially identical (Parˇ enicova´ et al., 2001). From morphological identifications based on the work of Raper and Fennell (1965), there is general agreement that Aspergillus aculeatus is the species commonly found in foods (Pitt and Hocking, 1997; Leong et al., 2004; Bau et al., 2005), so A. aculeatus is given priority here. If the two species are ever formally combined, Aspergillus japonicus has priority as it was described much earlier. Physiology. At 0.97 aw Aspergillus aculeatus grew at 20, 30 and 408C. At a lower water activity (0.92), it grew better at 308C than at 208C, and no growth was recorded at water activity values below 0.92 (0.87 or 0.82 aw) (Valero et al., 2007a). It has a high pectinolytic and cellulolytic enzyme activity (Adisa, 1989), and a fructosyltransferase with thermal and pH stability has recently been characterised (Ghazi et al., 2007). Mycotoxins. This species produces secalonic acid D (Frisvad et al., 2006a) and some other minor compounds (Frisvad and Samson, 1991). Secalonic acid D has significant animal toxicity (Ciegler et al., 1980), but a role in human or animal disease, especially from this species, has not been shown. Ecology. Aspergillus aculeatus causes a postharvest dry rot of tomatoes (Fajola, 1979). It is

297

commonly associated with grapes (Leong et al., 2004; Bau et al., 2005; Martı´ nez-Culebras and ´ 2007; Valero et al., 2007a) and is one of Ramon, the causes of bunch rot (Jarvis and Traquair, 1984). We isolated it in low numbers from Philippine and Indonesian peanuts and soybeans from the Philippines (Pitt et al., 1998a and unpublished). Additional species. Aspergillus japonicus Saito closely resembles A. aculeatus (see Taxonomy). A. japonicus sometimes produces larger colonies on CYA at 378C, up to 50 mm diam. Vesicles are smaller, usually less than 30 mm diam, and conidia are spherical. It does not produce secalonic acid D (Frisvad and Samson, 1991). The two species would be expected to have a similar physiology. Aspergillus japonicus has been isolated from Nigerian maize (Aja-Nwachukwu and Emejuaiwe, 1994). References. Raper and Fennell (1965); AlMusallam (1980), both as A. japonicus var. aculeatus and A. japonicus var. japonicus; Klich (2002), A. japonicus only.

Aspergillus candidus Link

Fig. 8.9

Colonies on CYA 15–20 mm diam, plane, low to moderately deep, dense, surface texture granular to floccose; mycelium white; conidial heads densely packed, persistently pure white to off-white; sometimes small amounts of clear exudate produced; reverse pale or yellow orange. Colonies on MEA 10–25 mm diam, similar to those on CYA except reverse dull brown. Colonies on G25N 10–16 mm diam, similar to those on CYA but conidial production often sparse and reverse pale or sometimes yellow. No growth at 58C. At 378C, colonies usually 20–25 mm diam, occasionally larger or absent, typically centrally umbonate and radially sulcate, velutinous or centrally floccose, with most characters similar to those on CYA at 258C. Conidiophores borne from surface or aerial hyphae, stipes usually 200–500(–1,000) mm long, with colourless, smooth to quite roughened walls; vesicles varying with isolate, 10–40 mm diam, bearing metulae and phialides over the entire surface, but numbers of metulae limited on the smallest heads; metulae variable, (5–)15–20(–30) mm long; phialides 5–9 mm long; conidia mostly spherical,

298

8 Aspergillus and Related Teleomorphs

Fig. 8.9 Aspergillus candidus (a) colonies on CYA and MEA, 7 days, 258C; (b) head, bar = 10 mm; (c) conidia, bar = 5 mm

2.5–3.5 mm diam, with smooth walls, borne in radiate heads. Distinctive features. The only species of Aspergillus with persistently white conidia, A. candidus is

readily distinguished from all species other than A. niveus. It differs from A. niveus by producing vesicles fertile over the entire area and metulae usually more than 10 mm long.

8.4

Genus Aspergillus

Physiology. The growth rates reported for this species vary considerably from optimum temperatures of 20–24 to 45–508C, minimum temperatures ranging from 3–4 to 11–138C and a maximum of 41–42 to 50–558C (see Pitt and Hocking, 1997). We have encountered a group of isolates from tropical dried fish, clearly identifiable as Aspergillus candidus, unable to grow at 378C. These figures are surprisingly conflicting for what appears to be a well defined species. The minimum aw for growth of Aspergillus candidus was reported as 0.75, by both Galloway (1935) and Ayerst (1969), after 14 days incubation at 258C and ca. 30 days at 308C, respectively. The optimum aw for growth was greater than 0.98 (Ayerst, 1969). Conidia of A. candidus showed 100% survival after heating for 10 min at 508C, but no survival after 10 min at 608C (Pitt and Christian, 1970). This species is more tolerant of low O2 than most Aspergillus species, being capable of growth in 0.45% O2. More than 15% CO2 was required to halve the growth rate in air (Magan and Lacey, 1984b). Tolerance to propionic acid used as a grain preservative is also high (Mu¨ller et al., 1981). Mycotoxins. A. candidus produces a range of secondary metabolites (Frisvad and Samson, 1991), but of these only kojic acid has even marginal toxicity. Ecology. Aspergillus candidus is widespread in foods. A xerophile, it is of common occurrence in stored cereals, where it may cause loss of germinability (Sinha and Wallace, 1977). It has been recorded from freshly harvested wheat, wheat during drying and in storage, cereal products, flour and refrigerated dough bran, bread, stored and mouldy maize (see Pitt and Hocking, 1997 and Lugauskas et al., 2006), milled rice (see Pitt and Hocking, 1997 and Taligoola et al., 2004), polished rice (Park et al., 2005a), yellow rice (Phillips et al., 1988), barley (see Pitt and Hocking, 1997 and Lugauskas et al., 2006) and cereal flakes (Weidenborner and Kunz, 1994). ¨ Aspergillus candidus is also a major species in nuts, including peanuts, hazelnuts and walnuts and pecans. A. candidus frequently occurs on salamis and other processed meats. Other sources include dried fish, cheesecake, soybeans, faba beans, sorghum, rapeseed and sunflower seed, mango powder, and health foods (see Pitt and Hocking, 1997).

299

Aspergillus candidus was present in many samples of commodities we examined from Southeast Asia, but usually at low levels (Pitt et al., 1993, 1994). The exception was Indonesian milled rice, where A. candidus was present in high levels (occasionally up to 100% infection) in 56% of samples (Pitt et al., 1998a). On the positive side, A. candidus has been recommended as a starter culture of low toxicity for processed meat manufacture (Grazia et al., 1986; Spotti et al., 1994). Aspergillus candidus has been reported from a wide range of human infections, including invasive aspergillosis and ear infections. It has also been isolated from birds (De Hoog et al., 2000). References. Raper and Fennell (1965); Domsch et al. (1980); Klich (2002).

Aspergillus carbonarius (Bainier) Thom Fig. 8.10 Aspergillus pulchellus (Speg.) Thom and Church Aspergillus fonsecaeus Thom and Raper

Colonies on CYA 60 mm or more diam, consisting of surface or subsurface white mycelium, surmounted by a layer of conidiophores; near margins individual conidial heads visible with the naked eye, more or less covering central areas, jet black; reverse colourless to pale yellow. Colonies on MEA 60 mm or more diam, similar to those on CYA but less dense; mycelium subsurface and inconspicuous; conidial heads usually individually visible over the entire colony, jet black; reverse uncoloured. Colonies on G25N 8–25 mm diam, of sparse white surface or subsurface mycelium surmounted by sparse black conidial heads; reverse uncoloured. No germination at 58C. At 378C, colonies 15–35 mm diam, of dense white to yellow mycelium; margins irregular; sporulation sparse, with visible individual heads, to very dense, jet black; reverse pale, pale brown, or grey to black. Conidiophores borne from surface or subsurface hyphae, 2.0–3.0 mm long, with heavy, smooth walls, uncoloured or grey; vesicles spherical or oblate, usually 60–90 mm diam, bearing closely packed metulae and phialides over the whole surface; metulae 12–18 mm long; phialides 9–12 (–15) mm long; conidia spherical or near, 6–7(–8) mm diam, black, with walls conspicuously roughened or sometimes spiky, borne in large, radiate heads.

300

8 Aspergillus and Related Teleomorphs

Fig. 8.10 Aspergillus carbonarius (a) colonies on CYA and MEA, 7 days, 258C; (b, c) heads, bars = 40 mm; (d) conidia, bar = 5 mm

Distinctive features. Aspergillus carbonarius is distinguished from related black Aspergillus species by producing conidia more than 6 mm in diameter. Colonies are jet black, and all heads produce metulae. Physiology. Aspergillus carbonarius can grow at 108C but not 7–88C, has an optimum near 308C and maximum near 418C (Bellı´ et al., 2004; Mitchell et al., 2004; Leong et al., 2006b; Marı´ n et al., 2006; Romero

et al., 2007). The optimum aw for growth is 0.96–0.98 (Mitchell et al., 2004; Leong et al., 2006b), with a minimum near 0.85 aw at 25–308C (Romero et al., 2007). No growth was observed at 0.82 aw in a glycerol based medium over the temperature range 15–358C (Romero et al., 2007). A. carbonarius grows over a wide pH range (2–10) (Esteban et al., 2005). Low levels (100 mg/l) of sodium metabisulphite, which is widely

8.4

Genus Aspergillus

used in the wine industry to treat grape juice before fermentation, stimulated growth of A. carbonarius, and high levels (720–1,000 mg/l) were required to inhibit growth by 90% (Pateraki et al., 2007). Aspergillus carbonarius is very tolerant of elevated levels of CO2. Compared with growth in air, 25% CO2 + 1% O2 stimulated growth, and an atmosphere of 50% CO2 + 1% O2 resulted in growth rates only slightly slower than the air control on grape juice medium at 0.985–0.93 aw (Pateraki et al., 2007). Conidia of A. carbonarius were more resistant to ultraviolet radiation than A. niger, but more sensitive than Alternaria alternata (Valero et al., 2007b). A. carbonarius conidia dried onto filter papers survived well at low aw (0.4 aw) and low temperature (18C), with little loss of viability after 618 days storage (Leong et al., 2006e). Mycotoxins. Ochratoxin A production by Aspergillus carbonarius was first reported by Horie (1995), Te´ren et al. (1996) and Wicklow et al. (1996). Since then, A. carbonarius has been recognised as the primary source of ochratoxin A contamination in grapes and grape products throughout the world (Battilani and Pietri, 2002; Caban˜es et al., 2002; Sage et al., 2002; Abarca et al., 2003; and many subsequent papers). A. carbonarius is also a potential source of ochratoxin A in coffee. It has been reported from coffee beans in Brazil (Taniwaki et al., 2003; Magnani et al., 2005), Ivory Coast (Kouadio et al., 2006), Uganda (Ngabirano et al., 2001), Thailand (Joosten et al., 2001) and Vietnam (Leong et al., 2007). Infection occurs postharvest, and ochratoxin A may be formed if beans are not dried efficiently (Taniwaki et al., 2003). Where grapes are further processed to make dried vine fruits, the black Aspergilli continue to grow, as in most producing countries grapes are dried in the sun without preservatives. Dried grapes (raisins, sultanas) can therefore readily contain unacceptable levels of ochratoxin A (Lombaert et al., 2004; Iamanaka et al., 2005; Jørgensen, 2005; Meyvaci et al., 2005). Aspergillus carbonarius produces ochratoxin A optimally at cooler temperatures: 158C and 0.95–0.97 aw or 208C and 0.98–0.99 aw. Little or no ochratoxin A is formed at temperatures above 358C. The lower aw limit for ochratoxin A production is near 0.92 (Mitchell et al., 2004; Esteban et al., 2004; Bellı´ et al., 2005; Marı´ n et al., 2006; Leong et al., 2006b). Ochratoxin A is produced over the pH

301

range 2–10 at 15 and 308C on CYA and YES media, respectively (Esteban et al., 2005). Sodium metabisulphite inhibits ochratoxin A production, but at high aw (0.985), at least 650–700 mg/l is required for 90% inhibition (Pateraki et al., 2007). Modified atmosphere storage (25 or 50% CO2) has little effect on ochratoxin formation, with 25% CO2 resulting in stimulation of toxin production (Pateraki et al., 2007). Application of fungicides to grapes can reduce fungal growth and ochratoxin production, but some fungicides appear to stimulate ochratoxin production (Bellı´ et al., 2006). Ochratoxin A is largely removed during the wine making process as it is bound to solid fractions such as wine lees and sediment. Use of some fining agents can also reduce ochratoxin levels in wine. Red wines retain slightly more ochratoxin A than white wines but, overall, the carryover from grapes into finished wine is between 1 and 8% (Leong et al., 2006c,d; Fernandes et al., 2007). Ecology. The primary food related habitat for A. carbonarius is grapes and the vineyard environment. A. carbonarius was first reported as an agent of Aspergillus bunch rot in India by Gupta (1956), well before the problem of ochratoxin A contamination of grape products was recognised. Since the late 1990s, there has been a concerted effort to understand the ecology of A. carbonarius and ochratoxin A formation in both wine grapes and dried grapes. A. carbonarius has been reported from grapes in Australia, South America and countries around the Mediterranean rim including Italy, Spain, Portugal, France, Greece, Turkey, Tunisia, Morocco, Lebanon and Israel (see Battilani et al., 2006 and references therein; El-Khoury et al., 2006). A. carbonarius can be isolated from vineyard soil and vine remnants (dried berries, bunch stems, dead canes, etc). It occurs on grapes from berry set to harvest, but populations increase from veraison onwards. A carbonarius is a saprophyte rather than a pathogen, so predisposing factors such as insect damage, infections with powdery mildew or Botrytis, or berry splitting due to rain or storm damage are needed to initiate Aspergillus bunch rot which generally occurs near harvest when sugar content of berries is highest (Leong et al., 2006a). A. carbonarius occurs on grapes during drying and can develop to high numbers on fruit that has been rain affected before harvest (Leong et al.,

302

8 Aspergillus and Related Teleomorphs

2004; 2006a). A. carbonarius has been reported from dried vine fruits from many countries including Spain (Abarca et al., 2003), Greece (Tjamos et al., 2004), Australia (Leong et al., 2004), Brazil (Iamanaka et al., 2005) and Argentina (Romero et al., 2005). Populations of Aspergillus carbonarius in vineyards can be reduced by vineyard management practices such as irrigation, pruning to improve air flow through vines, use of cover crops between rows and appropriate fungicide applications (Pollastro et al., 2005; Bellı´ et al., 2006; Leong et al., 2006a). Biocontrol of black Aspergilli on grapes using epiphytic yeasts has been proposed (Bleve et al., 2006). Other reported sources of Aspergillus carbonarius are figs (Doster et al., 1996), peanuts and maize (Magnoli et al., 2006a, b), bee pollen (Gonzalez et al., 2005), paprika (Almela et al., 2007), red bay berries (Li et al., 2002) and fermenting cocoa beans (Mounjouenpou et al., 2008). In our laboratory we have isolated A. carbonarius from dried vine fruits, walnuts and margarine. A. carbonarius isolated from Greek tannery wastes reduced tannins by 78%, significantly reducing the BOD of the effluent (Marakis, 1995). References. Raper and Fennell (1965); Klich (2002).

Aspergillus clavatus Desm.

Fig. 8.11

Colonies on CYA and MEA 30–45 mm diam, plane, of sparse surface mycelium surmounted by regular or irregular clusters of positively phototropic conidiophores up to 3 mm long, growing vertically if incubated in darkness; mycelium white; conidiophores readily visible under the stereomicroscope, with stout stipes and heads like match heads, with spore chains coloured greyish turquoise; clear exudate sometimes present in minute droplets; sometimes faint brown soluble pigment produced; reverse pale. Colonies on G25N 8–12 mm diam, of sparse, floccose, white mycelium and small, scattered conidiophores; reverse pale. No growth at 58C. At 378C, colonies 10– 30 mm diam, of white mycelium, with small blue grey heads; sometimes yellow brown soluble pigment produced; reverse yellow green. Conidiophores borne from subsurface or surface hyphae, stipes 1.5–3.0 mm long, with thick, smooth walls; vesicles narrow ellipsoids up to 250  70 mm, fertile over the whole area, bearing phialides only; phialides very closely packed, mostly 7–8 mm long; conidia ellipsoidal, 3.0–4.5 mm long, smooth walled,

borne in radiate heads, each splitting into two or more ordered columns in age. Distinctive features. Long ellipsoidal vesicles, heads with phialides only and grey blue conidia set Aspergillus clavatus apart from other species. Taxonomy. The relationship of Aspergillus clavatus with closely related species was studied by Varga et al. (2003). Physiology. Panasenko (1967) reported that Aspergillus clavatus had an optimal growth temperature near 258C, a minimum of 5–68C and a maximum of 428C. Similar figures were reported by Northolt et al. (1978). The minimum aw permitting growth was reported as 0.88 aw by Panasenko (1967) and graphed as near 0.87 aw by Northolt et al. (1978). Mycotoxins. This species produces patulin which has the potential to cause ill-thrift in animals and may have harmful effects in man. Patulin is discussed under Penicillium expansum. Aspergillus clavatus produces a number of other mycotoxins of marginal importance including kojic acid, cytochalasin E, tremorgenic tryptoquivalines and glyantrypine (Varga et al., 2003; Sabater-Vilar et al., 2004). Patulin was produced by Aspergillus clavatus at 0.99 aw but not 0.95 aw on laboratory media (Northolt et al., 1978). Ecology. A. clavatus is of particularly common occurrence in barley during malting and can build to unacceptably high levels if malting temperatures are elevated or spontaneous heating occurs (Flannigan et al., 1984; Flannigan, 1986; SabaterVilar et al., 2004). In extreme cases blue green mats of A. clavatus may form on grain during malting (Shlosberg et al., 1991). Under these conditions it can be allergenic and is reported to be the cause of ‘‘malt workers’ lung’’ (Riddle et al., 1968; Flannigan, 1986; Sabater-Vilar et al., 2004). Aspergillus clavatus is mostly associated with cereals and has been reported from wheat, flour, bread, rice and maize (see Pitt and Hocking, 1997). Other sources include red peppers, pecan nuts, health foods, biltong and salted dried fish (see Pitt and Hocking, 1997). In Southeast Asian food commodities, Aspergillus clavatus occurred at low levels in maize from Thailand and the Philippines; peanuts, kemiri nuts and pepper from Indonesia; and milled rice, soybeans and mung beans from the Philippines. Levels were slightly higher in copra from Thailand (Pitt et al., 1993, 1994, 1998a and unpublished).

8.4

Genus Aspergillus

303

Fig. 8.11 Aspergillus clavatus (a) colonies on CYA and MEA, 7 days, 258C; (b, c) heads, bars = 25 mm; (d) conidia, bar = 5 mm

References. Raper and Fennell (1965); Domsch et al. (1980); Klich (2002).

Aspergillus flavipes (Bainier & Sartory) Thom & Church Fig. 8.12 Teleomorph: Fennellia flavipes B.J. Wiley & E.G. Simmons Colonies on CYA 15–30 mm diam, deep but velutinous, lightly radially sulcate, occasionally floccose

centrally; mycelium white to yellow; conidial heads borne uniformly over the colony surface, pale greyish orange; clear exudate sometimes present; reverse pale, pale yellow or light to yellowish brown. Colonies on MEA 20–35 mm diam, plane, velutinous or sometimes slightly granular; mycelium white, inconspicuous; conidial production light, pale orange brown; reverse greyish yellow to golden brown. Colonies on G25N 8–12 mm diam, plane, dense to floccose, white to greyish orange; reverse brown. No growth at 58C. At 378C, colonies 5–20 mm diam.

304

8 Aspergillus and Related Teleomorphs

Fig. 8.12 Aspergillus flavipes (a) colonies on CYA and MEA, 7 days, 258C; (b, c, d) heads, bars = 10 mm; (e) conidia, bar = 5 mm

Cleistothecia fide Wiley and Simmons (1973) yellow, embedded in discrete clumps of rounded to elongate Hu¨lle cells; ascospores maturing in about 3 weeks, hyaline, subspheroidal, 6.4–8.0  5.6–6.4 mm, smooth walled, with an inconspicuous longitudinal groove. Conidiophores borne from surface or aerial hyphae, stipes 150–600 mm long, uncoloured to pale brown, walls smooth to slightly roughened, broadening gradually into spathulate to subspheroidal vesicles mostly 10–20 mm diam, bearing metulae and phialides over the upper half to two thirds; metulae and phialides 5–7 mm long; conidia spherical, 2.0–3.0 mm diam, smooth walled, borne in radiate to loosely columnar heads.

Distinctive features. This species is distinguished by pale greyish orange colonies with metulae and phialides bearing small, smooth walled, spherical conidia. Taxonomy. Aspergillus flavipes occasionally produces a teleomorph, described by Wiley and Fennell (1973) and transferred to Fennellia by Wiley and Simmons (1973). Physiology. This species grows from 6–7 to 38–408C, with an optimum near 26–288C (Domsch et al., 1980). It is capable of growth in at least 25% (w/v) NaCl, corresponding to 0.82 aw (Tresner and Hayes, 1971).

8.4

Genus Aspergillus

Mycotoxins. This species produces a variety of secondary metabolites (Frisvad and Samson, 1991) including lovastatin (Valera et al., 2005), but no known mycotoxins. Ecology. Aspergillus flavipes is common in tropical soils (Domsch et al., 1980), but uncommon in foods. It has been reported to cause spoilage of a variety of tropical fruit and cassava. It has been reported in wheat from France and we isolated it at low levels from cassava, peanuts and coriander in Southeast Asia (Pitt and Hocking, 1997).

305

References. Raper and Fennell (1965); Wiley and Fennell (1973); Domsch et al. (1980); Klich (2002).

Aspergillus flavus Link

Fig 8.13

Colonies on CYA 60–70 mm diam, plane, sparse to moderately dense, velutinous in marginal areas at least, often floccose centrally, sometimes deeply so; mycelium only conspicuous in floccose areas, white; conidial heads usually borne uniformly over the whole colony, but sparse or absent in areas of

Fig. 8.13 Aspergillus flavus (a) colonies on CYA and MEA, 7 days, 258C; (b, c) heads, bars = 20 mm; (d) conidia, bar = 5 mm

306

floccose growth or sclerotial production, characteristically greyish green, yellow green or olive yellow, but sometimes pure yellow, then becoming greenish in age; sclerotia produced by about 50% of isolates, at first white, becoming deep reddish brown, density varying from inconspicuous to dominating colony appearance and almost entirely suppressing conidial production; exudate sometimes produced, clear, or reddish brown near sclerotia; reverse uncoloured or brown to reddish brown beneath sclerotia. Colonies on MEA 50–70 mm diam, similar to those on CYA although usually less dense. Colonies on G25N 25–40 mm diam, similar to those on CYA or more deeply floccose and with little conidial production, reverse pale to orange or salmon. No growth at 58C. At 378C, colonies usually 55–65 mm diam, similar to those on CYA at 258C, but more velutinous, with olive conidia, and sometimes with more abundant sclerotia. Sclerotia produced by some isolates, at first white, rapidly becoming hard and reddish brown to black, spherical, usually 400–800 mm diam, but less than 400 mm in ‘‘S’’ strains (Cotty, 1989). Teleomorph not known. Conidiophores borne from subsurface or surface hyphae, stipes 400 mm to 1 mm or more long, colourless or pale brown, rough walled; vesicles spherical, 20–50 mm diam, fertile over three quarters of the surface, typically bearing both metulae and phialides, but in some isolates a proportion or even a majority of heads with phialides alone; metulae and phialides of similar size, 7–10 mm long; conidia spherical to subspheroidal, usually 3.5–5.0 mm diam, with relatively thin walls, finely roughened or, rarely, smooth. Distinctive features. Aspergillus flavus and A. parasiticus are distinguished by their rapid growth at both 25 and 378C and their bright yellow green (or less commonly yellow) conidial colours. The definitive difference between the two species is that A. flavus produces conidia which are rather variable in shape and size, have relatively thin walls and range from smooth to moderately rough, the majority being finely rough. In contrast, conidia of A. parasiticus are spherical and have relatively thick, rough walls. In addition, vesicles of A. flavus are larger, up to 50 mm in diameter, and usually bear metulae, while vesicles of A. parasiticus rarely exceed 30 mm in diameter and metulae are uncommon.

8 Aspergillus and Related Teleomorphs

Taxonomy. Klich and Pitt (1985, 1988) showed that metulae production, the principal character used by Raper and Fennell (1965) for distinguishing Aspergillus flavus from A. parasiticus, was not a satisfactory criterion. Some A. flavus isolates produce very few metulae. Careful study showed several differences between the two species, the most useful of which are mentioned above. The criteria used here provide a 100% correlation with mycotoxin production (see below). Kurtzman et al. (1986b) reduced Aspergillus parasiticus to the status of subspecies, as A. flavus subspecies parasiticus (Speare) Kurtzman et al. The domesticated species A. oryzae and A. sojae were reduced to the status of variety. Kurtzman et al. (1986b) based these changes on a DNA hybridisation study using only a single strain from each species. Apart from doubt over the validity of this work, or the meaning of DNA hybridisation percentages in these fungi, compelling reasons exist for maintaining distinct species names for these taxa, as has been done here. First, A. flavus differs reliably from A. parasiticus in the morphological features noted above. Second, some isolates of A. flavus produce B aflatoxins and some form cyclopiazonic acid, whereas A. parasiticus isolates all produce both B and G aflatoxins and do not form cyclopiazonic acid. These differences are of both scientific and commercial importance. Third, much evidence suggests that A. parasiticus has a more limited geographical range than A. flavus. Fourth, while A. flavus and A. parasiticus are both recognised as producers of carcinogenic mycotoxins, A. oryzae and A. sojae are widely used in food fermentations. It is of great practical importance to the food industry and to regulatory authorities everywhere that the naturally occurring highly toxigenic species have names quite distinct from those used in food fermentations. On this latter ground alone, maintenance of all four species names is a matter of necessity. All four species names are protected in the NCU list (Pitt and Samson, 1993). Physiology. A problem in assessing physiological data for Aspergillus flavus and related species has been inaccurate identification. Most physiological studies reported on A. flavus up to 1990 in fact dealt with A. parasiticus or A. nomius (Pitt, 1993b). Fortunately, differences among these species do not appear to be great, but some data below cannot

8.4

Genus Aspergillus

with certainty be identified as relating to A. flavus. Reported growth temperatures for A. flavus show some variation: a minimum near 10–128C, a maximum near 43–488C and an optimum near 338C appear to be most frequently mentioned (see Pitt and Hocking, 1997). Differences in water activities permitting growth have also been rather variable: from a low of 0.78 aw at 338C (Ayerst, 1969) to 0.84 at 258C (Pitt and Hocking, 1977). Pitt and Miscamble (1995) reported a minimum of 0.82 at 258C, 0.81 at 308C and 0.80 at 378C. Data from that paper were used to provide a predictive model for A. flavus growth in relation to aw and temperature (Gibson et al., 1994). High temperatures (418C) decreased germination rates of A. flavus by 45% compared with germination at 308C (Araujo and Rodrigues, 2004). Growth of A. flavus occurred over the pH range 2.1–11.2 (the entire range examined) at 25, 30 and 378C, with optimal growth over a broad range from pH 3.4–10 (Wheeler et al., 1991) and a peak near 7.5 (Olutiola, 1976). The use of acetic acid to adjust the pH of growth media to 4.5 reduced the germination of A. flavus conidia, whereas lactic and hydrochloric acids had no effect on germination (Araujo and Rodrigues, 2004). The heat resistance of A. flavus has been studied under various conditions by several authors. The most reliable figures indicate a D45 value of more than 160 h, a D50 of 16 h, a D52 of 40–45 min and a D60 of 1 min, at neutral pH and high aw, with z values from 3.3 to 4.1C8 (Beuchat, 1981b,c; Doyle and Marth, 1975a; ICMSF, 1996a). The addition of phosphine, used to control insects, to grain at 0.80 or 0.86 aw reduced growth of Aspergillus flavus while having little effect on the survival of conidia (Hocking and Banks, 1991). After 42–48 h incubation on AFPA (Pitt et al., 1983), colonies of Aspergillus flavus and A. parasiticus exhibit a brilliant orange yellow reverse colouration (see Chapter 4). Mycotoxins. Aspergillus flavus is the main source of aflatoxins, the most important mycotoxins in the world’s food supplies. Aflatoxins are produced in nature by A. flavus, A. parasiticus and a number of other species, including A. nomius, which are of little practical importance in foods. The four major naturally produced aflatoxins are known as aflatoxins B1, B2, G1 and G2. ‘‘B’’ and ‘‘G’’ refer to the blue and green fluorescent colours, respectively, produced

307

under UV light on thin layer chromatography plates, while the subscript numbers 1 and 2 indicate major and minor compounds, respectively. Aflatoxins are both acutely and chronically toxic to animals, including man. They have long been know to produce four distinct effects: acute liver damage, liver cirrhosis, induction of tumours and teratogenic effects (Stoloff, 1977). However more recent information indicates that the consequences of prolonged aflatoxin exposure are more widespread, including immunosuppression and interference with protein uptake (Williams et al., 2004). Outbreaks of acute aflatoxin poisoning are not common, but are a recurrent public health problem. A 1974 outbreak of hepatitis that affected 400 Indian people, of whom 100 died, was traced to maize heavily contaminated with Aspergillus flavus and containing up to 15 mg/kg of aflatoxins (Krishnamachari et al., 1975). Consumption of toxins by some of the affected adults was calculated to be 2–6 mg in a single day. It can be concluded that the acute lethal dose for adult humans is of the order of 10 mg. In 2004, one of the largest and most severe outbreaks of aflatoxicosis occurred in eastern Kenya, where 317 people fell ill and 125 died. This was followed by a second outbreak in 2005 during which a further 25 people died (Strosnider et al., 2006). A case study conducted soon after the first outbreak revealed that mouldy maize was the source of aflatoxin contamination (Azziz-Baumgartner et al., 2005). Aflatoxin B1 (AFB1) has been demonstrated, in a variety of animal species, to be the most potent liver carcinogen known. Aflatoxin B1 has been linked to human hepatocellular carcinoma in several regions of Africa and Southeast Asia (Wogan, 1992; Wang and Groopman, 1999). In consequence, AFB1 is regarded as a category I human carcinogen by the International Agency for Research on Cancer (IARC, 1993; Rastogi et al., 2006). Kuiper-Goodman (1991) concluded that between 5 and 9 mg/kg body weight/day of AFB1 intake had a 50% probability of increasing the incidence of primary liver cancer in a population by a factor of 1 case per 10,000 population. Based on those figures and aflatoxin levels in Indonesian diets (Pitt and Hocking, 1996), Lubulwa and Davis (1994) estimated that the minimum numbers of deaths in Indonesia from aflatoxins in peanuts and maize exceeded 22,000 per annum.

308

A strong synergy is observed between aflatoxin and hepatitis B (HBV) virus in the aetiology of liver cancer (Williams et al., 2004). The relative risk of cancer for HBV antigen positive patients rises from about 5 to about 60 when combined with aflatoxin exposure. In some areas where HBV and aflatoxin co-occur, hepatomas are the predominant cancer and may be a predominant cause of death (Williams et al., 2004). This known carcinogenicity is so high that every effort must be made to monitor levels in human foods and to reduce them as far as possible. When aflatoxin B1 and B2 are ingested by lactating cows, a proportion (ca. 1.5%; Frobish et al., 1986) is hydroxylated and excreted in the milk as aflatoxins M1 and M2, compounds of lower toxicity than the parent molecules. Children consume large quantities of milk and are also susceptible to the long term ill effects of aflatoxins, so concern soon mounted about the possible presence of aflatoxin M1 in milk, and studies in many countries reported positives (van Egmond, 1989). As a result stringent limits have been imposed on aflatoxin levels in feeds for dairy animals (van Egmond, 1989). A recent survey of UHT and pasteurised milk in Brazil indicated a high incidence (80%) of aflatoxin M1 in samples analysed; however, the concentration of aflatoxin M1 was below the 500 ng/l permitted limit (Garrido et al., 2003). Residues in other animal products are generally low. In particular, ruminant animals detoxify aflatoxins quite efficiently. Levels in eggs and meat from monogastric animals may be higher, but in general toxicity to the animal limits feeding of excessively contaminated feed, so minimising residues (Stoloff, 1977). It is believed that constant exposure to aflatoxins through contaminated foods leads to chronic diseases which also have a considerable impact on human health (Williams et al., 2004). Kwashiorkor, also known as protein energy malnutrition, is a leading cause of disease and death in children under 5 years of age in Northern Africa and elsewhere and it was Hendrickse et al. (1982) who first proposed a link with aflatoxin intake. Kwashiorkor was among the 10 most common admissions to one paediatric ward in Tanzania and had the third highest mortality rate (43%) after AIDS and septicaemia (Immerzeel et al.,

8 Aspergillus and Related Teleomorphs

2006). It is now established that chronic exposure to aflatoxins is correlated with the disease, in which hepatic metabolism is impaired (Fuhrman et al., 2004; Mu¨ller and Krawinkel, 2005). Hatem et al. (2005) reported the presence of aflatoxins in 80% of sera from infants with kwashiorkor. Aflatoxins have also been reported to be potent immunotoxic agent in both animals and humans and may be responsible for greatly increasing susceptibility to many kinds of disease agents in countries where aflatoxin ingestion is common (Wogan, 1992; Wang and Groopman, 1999; Williams et al., 2004). Cyclopiazonic acid (CPA) is produced by some strains of Aspergillus flavus. Its role in human health remains unclear, but evidence for its involvement in animal disease is strong (Cole, 1986). Occurrence at low levels in maize has been reported (Lee and Hagler, 1991; Giorni et al., 2007). The universal occurrence of Aspergillus flavus and aflatoxins in commodities such as peanuts, maize and cottonseed has prompted a great deal of work on causes, course and prevention, work too voluminous to report in detail here (for reviews see Diener et al., 1987; Payne, 1992). Earlier studies assumed that invasion was primarily a function of inadequate drying or improper storage, but more recent work has shown that, at least in developed countries, invasion before harvest is more important and is dependent primarily on plant stress induced by drought and/or high soil temperatures (Sanders et al., 1981; Cole et al., 1982). The problem can be overcome most effectively by irrigation, but unfortunately this is not a practical solution in many peanut growing regions. Preharvest invasion also occurs in maize, cottonseed, pistachios and figs (see Pitt and Hocking, 1997). In peanuts, reduction in aflatoxin levels during processing is accomplished by colour sorting of individual kernels after shelling. The process was developed originally to reject commercially unacceptable discoloured nuts, regardless of cause: but as fungal growth is a prime cause of discolouration, the process is also an effective nondestructive means of removing most aflatoxin-containing nuts. Maize and fig samples are screened for the presence of aflatoxin by the examination of cracked kernels or fruit by ultraviolet light (see Pitt and Hocking, 1997). No effective nonchemical testing techniques

8.4

Genus Aspergillus

exist for cottonseed or pistachios and, as with other commodities, nondestructive chemical assays are not available. The use of biological control, based on competitive exclusion, has been developed for peanuts and cottonseed (Pitt and Hocking, 2006; Dorner and Horn, 2007). This involves establishing high populations of nontoxigenic strains of Aspergillus flavus or A. parasiticus in the soil which then compete with naturally occurring aflatoxigenic strains, particularly when crops become stressed and susceptible to invasion. Dorner and Horn (2007) reported that treatment with a nontoxigenic A. flavus strain alone was more effective than a single strain of A. parasiticus and equally effective as a mixture of both strains in controlling toxigenic A. flavus in peanut crops. Counts of Aspergillus flavus in spices are often high, and concern has been expressed over the possible occurrence of aflatoxins in these commodities. Although the quantities of spices used in most foods are small, limiting any direct hazard, spices are used in the preparation of many types of foods, so high levels of A. flavus in spices are undesirable. Sterilisation of them by chemicals or irradiation is now widely practised. The presence of A. flavus in spices such as pepper may result in infections in hospital patients (De Bock et al., 1989). It has been suggested that the presence of aflatoxins in Aspergillus flavus conidia is a cause for concern. Individual spores can contain high concentrations: up to 1,000 mg/kg, an incredibly high concentration in comparison with foods (Wicklow and Shotwell, 1983). However, the minute weight of individual spores means that actual intake levels from spores are very low. Aflatoxins are readily absorbed into the body from spores ingested into the lungs, with detrimental effects on alveolar macrophage function (Miller, 1994b; Williams et al., 2004). Aflatoxins can also be absorbed through the skin. Experiments in mice showed that continued dermal exposure to AFB1, even at low doses, might induce tumors (Rastogi et al., 2006). Foods prepared from maize as grits or cakes, etc., such as are consumed in Southeastern United States and other areas, often contain significant aflatoxin levels (Stoloff and Friedman, 1976). In a recent survey of breakfast and infant cereals in Canada, 50% of samples contained detectable levels of aflatoxins. However, only 4% of the breakfast

309

samples and 1% of infant cereals had AFB1 levels exceeding the 0.1 mg/kg maximum limit set by the European Union (Tam et al., 2006). Flour (made from seed, pulses or cereal) and starch samples purchased from traditional bazaars and markets in Turkey showed that more than 60% of samples contained aflatoxins (Baydar et al., 2005). Samples of food products purchased from Japanese retail outlets containing maize, peanuts, buckwheat (flour and noodles), rice or sesame oil did not contain aflatoxins. However, 10 of the 21 samples of peanut butter tested were contaminated with high levels of AFB1 (Sugita-Konishi et al., 2006). From the few studies of the effects of physical factors on aflatoxin production, it appears that Aspergillus flavus can produce aflatoxins over the range 13–378C, and above 0.82 aw, with optima perhaps 16–318C and 0.95–0.99 aw (ICMSF, 1996a). Aflatoxin production by A. flavus in agricultural soils appears to be correlated with latitude, aflatoxigenic strains being more prevalent in the southern peanut growing regions of the United States. It appears that dry, hot conditions favour the presence of toxigenic strains of A. flavus (Horn, 2003). We have made similar observations: A. flavus strains from New Zealand produce aflatoxins less frequently and at lower concentrations than strains from Australia or Southeast Asia (our unpublished data). The influence of crop species has also been raised as a factor in selecting for toxigenic A. flavus in agricultural soils (Horn, 2003). Additional studies are still needed to provide a more accurate picture. Ecology. The quest for knowledge about potential aflatoxin problems means that Aspergillus flavus has been sought in every conceivable kind of foodstuff. A. flavus has become the most widely reported foodborne fungus, reflecting its economic importance and relative ease of recognition as much as its ubiquity. It is especially abundant in the tropics, and it has a particular affinity for nuts and oilseeds as substrates. Aspergillus flavus was prevalent in 97% of peanuts examined in Southeast Asia. Maize samples from Indonesia and the Philippines also contained high levels of A. flavus contamination. Other Southeast Asian commodities found to harbour A. flavus included kemiri nuts, peppercorns, velvet and talo beans, copra, cassava, sorghum, paddy rice and soybeans (see Pitt and Hocking, 1997).

310

Aspergillus flavus occurs in most types of nuts from time to time and is capable of causing spoilage and/or producing aflatoxins (see Pitt and Hocking, 1997). Reports include tree nuts, pistachios (Bayman et al., 2002a), halva containing pistachios (Var et al., 2007), hazelnuts and walnuts (Bayman et al., 2002a; Gu¨rses, 2006), coconut (Onifade and Jeff-Agboola, 2003), copra (Srinivasulu et al., 2003), pecans, and kola nuts (see Pitt and Hocking, 1997). Cereals are also a common source of Aspergillus flavus. Maize and maize products are a particular problem (see Pitt and Hocking, 1997; Sekiyama et al., 2005; Garcı´ a and Heredia, 2006; Giorni et al., 2007). Insects may be involved in infection (Garcı´ a and Heredia, 2006) and a number of practices including good storage conditions (Saleemullah et al., 2006) and planting genetically modified crops (Marasas and Vismer, 2003) are now employed to control aflatoxin levels. Contamination has also been reported commonly in other cereals and cereal products: wheat (Lugauskas et al., 2006; Giray et al., 2007); wheat flour (Kumar et al., 2002; Ogundare and Adetuyi, 2003) and flour products including bread (Lugauskas et al., 2006), pasta (Halt et al., 2004) and bran; barley (Lugauskas et al., 2006; Medina et al., 2006; see also Pitt and Hocking, 1997); paddy, milled and parboiled rice (see Pitt and Hocking, 1997); rice and rice bran (Sales and Yoshizawa, 2005a); sorghum (da Silva et al., 2004; Lugauskas et al., 2006); and pearl millet (Wilson et al., 2006). However, unlike the situation with crops high in oil, spoilage by A. flavus or aflatoxin production in small grain cereals is almost always the result of poor handling. Aflatoxin levels in properly handled small grains are usually negligible (Stoloff, 1977; Pohland and Wood, 1987). Spices of many kinds frequently contain Aspergillus flavus (Elshafie et al., 2002; Gatti et al., 2003; Mandeel, 2005 and also see Pitt and Hocking, 1997). Ionising radiation (see Pitt and Hocking, 1997) and treatment of fresh plant materials before grinding or immediately after processing can reduce fungal and bacterial loads and aflatoxin levels in spices (Schweiggert et al., 2005). Aspergillus flavus is common in green coffee beans (Martins et al., 2003; Magnani et al., 2005) and herbal drugs (Rizzo et al., 2004). Other sources of A. flavus include chickpeas (Singh et al., 2005),

8 Aspergillus and Related Teleomorphs

pigeon peas (Reddy et al., 2006), soybeans (Ahammed et al., 2006), olives, rapeseed, amaranth seeds (see Pitt and Hocking, 1997), mustard seeds (Ahmad and Sinha, 2006), sesame seeds (Singh et al., 2003; Pillai et al., 2003); freshly harvested and stored blackgram (Goyal and Jain, 1998), sunflower seeds (Narasimhan and Muthumary, 2005) and betel nuts (see Pitt and Hocking, 1997). Processed and smoked meats, bacon (see Pitt and Hocking, 1997), milk (Garrido et al., 2003; Ruggia and Galiero, 2005) and cheese (Hayaloglu and Kirbag, 2007) often contain A. flavus and sometimes aflatoxins. Some processed fish products from Nigeria and Sierra Leone supported growth of A. flavus but it was relatively uncommon in salted dried fish from Southeast Asia or dry cured hams in Spain (see Pitt and Hocking, 1997). Aspergillus flavus is capable of causing spoilage of some kinds of fresh fruit and vegetables, including citrus, tomatoes, peppers, litchis, pineapples and pomegranates, but it is not usually of great importance (Snowdon, 1990, 1991). It has recently been reported to cause rot of peaches in Greece (Michailides and Thomidis, 2007). Pathogenicity. Aspergillus flavus is one of the main agents of human allergic bronchial aspergillosis and of pulmonary infections in immunocompromised patients. It also causes ear infections (De Hoog et al., 2000). Additional species. Aspergillus oryzae (Ahlburg) Cohn is closely related to A. flavus and produces colonies of similar or slightly smaller size on the standard media. It is usually distinguishable from A. flavus on the standard media after 7 days only by a more floccose and lightly sporing appearance and sometimes a tendency towards pale brown conidial colours. Colonies of A. oryzae change in conidial colour from green towards olive brown with continued incubation at 258C for 7–14 days. Colonies of A. flavus and A. parasiticus remain yellow green or become greyish green under these conditions. Conidial heads of A. oryzae usually bear metulae and phialides, and conidia are usually larger than those of A. flavus, with thin, smooth to finely roughened walls. A. oryzae can be distinguished from A. flavus by amplified fragment length polymorphism (AFLP) (Monteil et al., 2003; Lee et al., 2004). Aspergillus oryzae is of great economic importance, as it forms the basis of much of the fermented

8.4

Genus Aspergillus

311

food industry in Japan and other parts of Asia (see Pitt and Hocking, 1997). Tane koji, prepared by growing A. oryzae on cooked rice, provides a source of enzymes used in the production of shoyu (soy sauce), miso, hamanatto and other important Oriental products, which are mostly used as food flavourings (Tanaka et al., 2006). Aspergillus oryzae is rarely isolated from sources other than fermented foods and it is reasonable to believe that it is a domesticated form of A. flavus, adapted by centuries of use in fermented food manufacture (Wicklow, 1983). Unlike A. flavus, A. oryzae is not known to produce aflatoxins. However, fermented foods may not be as free from the hazards of mycotoxins as is popularly believed as strains of A. oryzae were found to produce cyclopiazonic acid and kojic acid (Tanaka et al., 2006). Additional species. Aspergillus nomius Kurtzman et al. (Kurtzman et al., 1986a) is also closely related to A. flavus. In the absence of sclerotia, the two species are morphologically indistinguishable. The major differences are that A. nomius produces both B and G aflatoxins and that sclerotia of A. nomius are bullet shaped, not spherical like those of A. flavus. Physiologically, A. nomius resembles A. flavus, but does not grow at quite such low water activities, with 0.83 aw the minimum at 25 and 308C and 0.81 at 378C (Pitt and Miscamble, 1995). Aspergillus nomius is a comparatively rare species, originally isolated from alkali bees in the United States (Kurtzman et al., 1986a). However, it has been reported from agricultural and native forest soils in Thailand where it was found to produce greater amounts of aflatoxins than A. flavus (Ehrlich et al., 2007), and we isolated it from a range of Thai commodities: maize, peanuts, soybeans, cassava and black beans, but always at low levels (Pitt et al., 1993, 1994). References. Raper and Fennell (1965); Domsch et al. (1980); Kurtzman et al. (1986a); Klich and Pitt (1988); Klich (2002).

Aspergillus fumigatus Fresen.

Fig. 8.14

Colonies on CYA 40–60 mm diam, plane or lightly wrinkled, low, dense and velutinous or with a sparse, floccose overgrowth; mycelium inconspicuous, white; conidial heads borne in a continuous, densely packed layer, greyish turquoise to dark

turquoise (blue green); clear exudate sometimes produced in small amounts; reverse pale or greenish. Colonies on MEA 40–60 mm diam, similar to those on CYA but less dense and with conidia in duller colours; reverse uncoloured or greyish. Colonies on G25N less than 10 mm diam, sometimes only germination, of white mycelium. No growth at 58C. At 378C, colonies covering the available area, i.e. a whole Petri dish in 2 days from a single point inoculum, of similar appearance to those on CYA at 258C, but with conidial columns longer and conidia darker, greenish grey to pure grey. Conidiophores borne from surface hyphae, stipes 200–400 mm long, sometimes sinuous, with colourless, thin, smooth walls, enlarging gradually into pyriform vesicles; vesicles 20–30 mm diam, fertile over half or more of the enlarged area, bearing phialides only, the lateral ones characteristically bent so that the tips are approximately parallel to the stipe axis; phialides crowded, 6–8 mm long; conidia spherical to subspheroidal, 2.5–3.0 mm diam, with finely roughened or spinose walls, forming radiate heads at first, then well defined columns of conidia. Distinctive features. This distinctive species can be recognised in the unopened Petri dish by its broad, velutinous, bluish colonies bearing characteristic, well defined columns of conidia. Growth at 378C is exceptionally rapid. Conidial heads are also diagnostic: pyriform vesicles bear crowded phialides which bend to be roughly parallel to the stipe axis. Care should be exercised in handling cultures of this species (see Chapter 4). Physiology. Undoubtedly the most important physiological character of Aspergillus fumigatus is its thermophilic nature: its minimum growth temperature is near 128C, the optimum 40–428C and maximum near 558C (Panasenko, 1967; Ayerst, 1969: Evans, 1971; Domsch et al., 1980). A. fumigatus is a marginal xerophile, Ayerst (1969) recording 0.82 aw as the minimum for growth, near 408C. It required more than 15% CO2 to reduce growth rates by half at 0.98 aw, but only 2.5% at 0.90 aw (Magan and Lacey, 1984b). Mycotoxins. Aspergillus fumigatus produces fumitremorgens, verruculogen and gliotoxin and a role in animal disease seems likely (Cole, 1981a,b; Dorner et al., 1984; Moreau, 1990; Frisvad et al., 2006a). Evidence suggests gliotoxin is important in

312

8 Aspergillus and Related Teleomorphs

Fig. 8.14 Aspergillus fumigatus (a) colonies on CYA and MEA, 7 days, 258C; (b, c) heads, bars = 10 mm; (d) conidia, bar = 5 mm

the invasion of animal and bird lungs by A. fumigatus (Richard et al., 1994). The presence of A. fumigatus in pepper used in foods for hospital patients is undesirable (De Bock et al., 1989). The fumigaclavines, produced by Aspergillus fumigatus, are related to ergot alkaloids (Panaccione and Coyle, 2005). A. fumigatus also produces fumagillin, a compound with some antibacterial and antifungal activity, which is also an angiogenesis inhibitor (Yang et al., 2003).

Ecology. The prime habitat for Aspergillus fumigatus is decaying vegetation, particularly grass clippings, in which it causes spontaneous heating (Cooney and Emerson, 1964). Major habitats are therefore in cocoa beans during and after fermentation and in spices (see Pitt and Hocking, 1997). It has been recorded frequently from stored commodities in the tropics, where its ability to grow simultaneously at low aw and at high temperatures provides an ecological advantage, e.g. stored oilseeds, stored eggs,

8.4

Genus Aspergillus

313

copra, soybeans and vegetables (see Pitt and Hocking, 1997). It has been reported as one cause of ‘‘Rio’’ off-flavours in coffee (Liardon et al., 1992), but it is not usually an important spoilage fungus. Cereals are a common source, e.g. wheat (Soldevilla et al., 2005; Lugauskas et al., 2006), rice, parboiled rice and barley (see Pitt and Hocking, 1997). It is quite common on cured and processed meats, especially in the tropics (Sabater-Vilar et al., 2003 and see also Pitt and Hocking, 1997). A. fumigatus also occurs on nuts, e.g. hazelnuts, walnuts, peanuts (see Pitt and Hocking, 1997) and cashews (Adebajo and Diyaolu, 2003). Other sources include dried fish, baladi bread, corn snacks, rootstock snacks, melon seeds, mango pickles, dried onion, low fat buffalo milk cottage cheese, processed cheese (see Pitt and Hocking, 1997) and kuflu cheese (Hayaloglu and Kirbag, 2007). A. fumigatus isolated from marine sediments produced gliotoxin under hypersaline conditions. After 6 days in gliotoxin contaminated seawater, fresh mussels were able to accumulate gliotoxin at levels up to 2.9 mm/mg in meat (Grovel et al., 2003). Pathogenicity. Aspergillus fumigatus is the main agent of pulmonary aspergillosis in man and birds. However, disseminated infections in patients without apparent immune disorders are extremely rare (De Hoog et al., 2000). References. Raper and Fennell (1965); Domsch et al. (1980); Klich (2002).

Aspergillus niger Tiegh. nom. cons.

Fig. 8.15

Colonies on CYA 60 mm or more diam, usually covering the whole Petri dish, plane, velutinous, of low, usually subsurface, white mycelium, surmounted by a layer of closely packed, dark brown to black conidial heads, ca. 2–3 mm high; reverse usually pale, sometimes pale to bright yellow. Colonies on MEA varying from 30 to 60 mm diam, usually smaller than those on CYA and often quite sparse by comparison, otherwise similar. Colonies on G25N 18–30 mm diam, plane, velutinous, with white or pale yellow mycelium visible at the margins, otherwise similar to those on CYA; reverse pale or occasionally with areas of deep brown. No growth at 58C. At 378C, colonies 60 mm or more diam, covering the available space, sometimes sulcate, otherwise similar to those on CYA at 258C.

Conidiophores borne from surface hyphae, 1.0–3.0 mm long, with heavy, hyaline, smooth walls; vesicles spherical, usually 50–75 mm diam, bearing closely packed metulae and phialides over the whole surface; metulae 10–15 mm long, or sometimes more; phialides 7–10 mm long; conidia spherical, 4–5 mm diam, brown, with walls conspicuously roughened or sometimes striate, borne in large, radiate heads. Distinctive features. One of the best known of all fungal species, Aspergillus niger is distinguished by its spherical black conidia, derived from colonies which show little or no other colouring. A. carbonarius (Bainier) Thom, which produces conidia 7–10 mm in diameter and A. awamori Nakaz., which produces finely roughened conidia, are closely related. A. awamori is used in food fermentations and is perhaps a domesticated form of A. niger. Taxonomy. The name Aspergillus niger, in use for most of this century, is predated by other valid names which have priority under the Botanical Code. A. niger was conserved by the International Botanical Congress in 1993 (Greuter et al., 1994). The formal wording ‘‘nom. cons.’’ after the name indicates this protected status. Molecular studies have shown that A. niger consists of two species, A. niger and A. tubingensis, separated at first by RFLP analysis (Kusters-van Sommeren et al., 1991; Varga et al., 1994) and later by sequence analysis (Peterson, 2000). The species are morphologically indistinguishable, and for practical purposes, the name A. niger can be used for both. Physiology. Growth temperatures for Aspergillus niger are minimum 6–88C, maximum 45–478C, and optimum 35–378C (Panasenko, 1967; Leong et al., 2006b). However, Palacios-Cabrera et al. (2005) reported no growth at 88C on three different media. A. niger is a xerophile: Ayerst (1966) reported germination at 0.77 aw at 358C. Only slight differences in growth rates were observed on media based on NaCl or glycerol, or of pH 4.0 and 6.5, at various water activities (Avari and Allsopp, 1983). A. niger is able to grow down to pH 2.0 at high aw (Pitt, 1981). Microwaves were found to have no effect on conidia of A. niger, apart from that from substrate heat (Ishitani et al., 1981). A. niger was relatively resistant to UV-C radiation but more sensitive than A. carbonarius or A. alternata (Valero et al., 2007b).

314

8 Aspergillus and Related Teleomorphs

Fig. 8.15 Aspergillus niger (a) colonies on CYA and MEA, 7 days, 258C; (b) head, bar = 15 mm; (c) heads, bar = 10 mm; (d) conidia, bar = 5 mm

Mycotoxins. Aspergillus niger has usually been regarded as a benign fungus and has been widely used in enzyme production and ingredients for food processing. It holds GRAS (generally regarded as safe) status from the US Food and Drug Administration. However, 2 of 19 A. niger isolates were reported to produce ochratoxin A by Abarca et al. (1994) and further research has shown than many more isolates of A. niger aggregate can produce ochratoxin A under various conditions (Bellı´ et al., 2004; Esteban et al. 2004, 2006 a,b; Leong et al., 2006b). Usually only a low percentage of A. niger

isolates are able to produce ochratoxin A. Frisvad and Samson (2004) reported that A. tubingensis does not produce ochratoxin A. However, recent studies have disagreed. Medina et al. (2005) and Perrone et al. (2006) have both reported the production of ochratoxin A by some isolates of A. tubingensis after identification by molecular methods, 3 of 21 (14%) and 5 of 20 (25%), respectively. Fumonisin B2 has recently been detected in cultures of three full genome sequenced strains of Aspergillus niger, including the ex type culture (Frisvad et al., 2007).

8.4

Genus Aspergillus

Ecology. Among the fungi most commonly reported from foods, Aspergillus niger is more prevalent in warmer climates, both in field situations and stored foods. The black spores provide protection from sunlight and UV light, providing a competitive advantage in such habitats (Valero et al., 2007b). A. niger is very frequently isolated from sun dried products, such as vine fruits (Abarca et al., 2003; Leong et al., 2004; Magnoli et al., 2004; Iamanaka et al., 2005), dried, smoked and cured fish, biltong, cocoa beans (see Pitt and Hocking, 1997) and spices (Elshafie et al., 2002; Mandeel, 2005). A similar physiology to A. flavus means that the two species compete in many situations. Various suggestions have been made for using A. niger as a competitor to A. flavus in the field (e.g. Wicklow et al., 1987; Shantha et al., 1990; Paster et al., 1992), but in our opinion A. niger is too destructive for this to be a practical proposition. Aspergillus niger is by far the most common Aspergillus species responsible for postharvest decay of fresh fruit, including apples, pears, peaches, citrus, grapes, figs, strawberries, mangoes and melons (Snowdon, 1990 and also see Pitt and Hocking, 1997). Most of these diseases are sporadic and of minor significance. A niger has also been reported to cause serious losses in tomatoes (Muhammad et al., 2004) and breadfruit (Omobuwajo and Wilcox, 1989) and in some vegetables, especially onions and garlic (Rath and Mohanty, 1985; Snowdon, 1991; Sinha et al., 1994), stored cabbages (Lugauskas et al., 2005) and yams (Adeniji, 1970a; Ogundana, 1972). It is frequently isolated from fresh grapes where it may be involved in Aspergillus bunch rot (Valero et al., 2005; Lasram et al., 2007; Fredj et al., 2007; Ponsone et al., 2007; Iamanaka et al., 2005). It is one species causing thread mould spoilage of cheese (Hocking and Faedo, 1992). Aspergillus niger is among the most common fungi isolated from nuts, especially peanuts, and also pecans (see Pitt and Hocking, 1997), pistachios, almonds and walnuts (Bayman et al., 2002a). It has been isolated from cashews (Adebajo and Diyaolu, 2003), kola nuts (Adebajo, 1994), coconut (Zohri and Saber, 1993) and copra (Srinivasulu et al., 2003). Cereals and oilseeds are also frequent sources, especially maize (see Pitt and Hocking, 1997; also Magnoli et al., 2007a for a recent review), corn snacks (Caldas et al., 2002) and also barley (see

315

Pitt and Hocking, 1997; Soldevilla et al., 2005; Lugauskas et al., 2006; Viswanath et al., 2006), soybeans (Aziz et al., 2006), rapeseed (Magan et al., 1993; Ahamad et al., 2003) and rape oil (Mondal and Nandi, 1984), sorghum (Navi et al., 2002 and see Pitt and Hocking, 1997), stored and parboiled rice (Aziz et al., 2006; Taligoola et al., 2004 and see Pitt and Hocking, 1997), blackgram (Ahmad, 1993), sunflower seeds (Begum et al., 2003), chickpeas (Ahmad and Singh, 1991) and pigeon peas (Maximay et al., 1992). Meat products are another common source (Farghaly et al., 2004; Hammad et al., 2006 and see Pitt and Hocking, 1997). Other records include cheese (Sinigaglia et al., 2004; Hayaloglu and Kirbag, 2007 and see Pitt and Hocking, 1997), olives (Roussos et al., 2006), fresh vegetables (Lugauskas et al., 2005) and strawberries, rootstock snacks and a variety of tropical products (see Pitt and Hocking, 1997). Aspergillus niger was commonly isolated during our study of Southeast Asian commodities. The highest contamination occurred in peanuts, maize and kemiri nuts. From Thailand, sorghum (58% of samples), cashews (53%) and copra (43%) contained A. niger, with overall infection rates of 7, 5 and 18%, respectively. Infections in pepper and spice from Indonesia and the Philippines were also quite high. Seventy eight per cent of coriander samples from Indonesia were infected with A. niger, with 18% infection overall. Infection rates with Aspergillus niger were lower in commodities other than the nuts and oilseeds mentioned above. Philippine soybeans were exceptional, with an infection in 47% of samples. Otherwise, soybeans from the other countries, mung beans, black beans, cowpeas, paddy rice and milled rice all were infected at no more than 2% of all particles examined (Pitt et al., 1993, 1994, 1998a and our unpublished data). References. Raper and Fennell (1965); Domsch et al. (1980); Al-Musallam (1980); Klich (2002).

Aspergillus niveus Blochwitz

Fig. 8.16

Teleomorph: Fennellia nivea (B.J. Wiley & E.G. Simmons) Samson Colonies on CYA 18–30 mm diam, plane or lightly sulcate, dense and velutinous; mycelium white,

316

8 Aspergillus and Related Teleomorphs

Fig. 8.16 Aspergillus niveus: (a) colonies on CYA and MEA, 7 days, 258C; (b, c, d) heads, bars = 10 mm; (e) conidia, bar = 5 mm

occasionally with bright yellow sectors; conidial heads closely packed, white to pale yellow; exudate clear, yellow brown or sometimes dull red; yellow soluble pigment sometimes present; reverse yellow brown to dark green. Colonies on MEA 18–30 mm diam, plane, similar to those on CYA but texture granular to floccose, exudate clear; reverse pale to dull yellow. Colonies on G25N 10–16 mm diam, dense to floccose, white; reverse pale to dull yellow. No growth at 58C. At 378C, colonies usually 30– 45 mm diam, similar to those on CYA at 258C, occasionally much smalle.

Cleistothecia fide Wiley and Fennell (1972) 80–130 mm diam, yellow, ascospores hyaline to yellow, 4.0–5.6 mm long, with an inconspicuous furrow, two low, longitudinal crests and spinose walls. Hu¨lle cells sometimes present on CYA, spherical to ellipsoidal, 10–18 mm long, yellow. Conidiophores borne from surface hyphae, stipes usually 100–500 mm long, with uncoloured, smooth walls; vesicles hemispherical to spathulate, 8–15 mm diam, bearing crowded metulae and phialides over the upper half to two thirds; metulae and phialides 5–9 mm long; conidia spherical to

8.4

Genus Aspergillus

317

subspheroidal, 2.5–3.5 mm diam, with smooth to finely roughened walls, borne in radiate to loosely columnar heads. Distinctive features. Aspergillus niveus is one of only two Aspergillus species with white or off-white conidia. It is distinguished from A. candidus by yellow conidial colours in age and more readily by vesicles which are fertile only over the upper half to two thirds, while those of A. candidus are fertile over the entire area. Taxonomy. This species occasionally produces a teleomorph, which was described as Aspergillus niveus by Wiley and Fennell (1973), was provided with the teleomorph name Emericella nivea by Wiley and Simmons (1973) and transferred to Fennellia by Samson (1979). Physiology. Aspergillus niveus grows strongly at higher temperatures, with a reported minimum of 11–138C, an optimum near 36–428C and a maximum at 47–488C (Domsch et al., 1980). Water relations have not been studied. Mycotoxins. A culture of Aspergillus niveus was toxic to ducklings (Scott, 1965). The toxicity of this species does not seem to have been further investigated, though Frisvad and Samson (1991) indicated that it may produce citrinin. Ecology. The main habitat of Aspergillus niveus is soil in the tropics (Domsch et al., 1980) and reports from foods are rare. However, we isolated this species from peanuts in Thailand, the Philippines and Indonesia; from soybeans, sorghum and black rice in Thailand; and milled rice in Indonesia (Pitt et al., 1993, 1994, and 1998a). Pectinase enzymes isolated from A. niveus were reported to improve the fermentation of tea leaves when compared with a commercial enzyme preparation (Angayarkanni et al., 2002). References. Raper and Fennell (1965); Wiley and Fennell (1973); Domsch et al. (1980), as Emericella nivea; Klich (2002).

Aspergillus ochraceus K. Wilh.

Fig. 8.17

Colonies on CYA 40–55 mm diam, plane or sulcate, low and velutinous or lightly floccose; mycelium white; conidial heads closely packed, light yellow to golden yellow; sclerotia sometimes produced, white when young, later pink to purple; clear exudate sometimes present, some exuded from stipe walls; reverse greyish orange to brown. Colonies

on MEA 40–55 mm diam, plane, similar to those on CYA but quite sparse; reverse pale yellow brown or slightly darker. Colonies on G25N 20–30 mm diam, plane, low and dense to deep and floccose, conidial production light to moderate, coloured as on CYA; reverse pale yellow or brown. No growth at 58C. At 378C, colonies up to 30 mm diam produced (Fig. 8.17). Conidiophores borne from surface hyphae, stipes 1.0–1.5 mm long, with yellowish to pale brown walls, finely to conspicuously roughened; vesicles spherical, 25–50 mm diam, bearing tightly packed metulae and phialides over the entire surface; metulae 15–20 mm long; phialides 9–12 mm long; conidia spherical to subspheroidal, 2.5–3.5 mm diam, with smooth to finely roughened walls, borne in radiate heads when young, splitting into two or more broad columns with maturity. Distinctive features. Aspergillus ochraceus produces yellow brown (ochre) conidia, borne on long stipes; vesicles bear metulae and phialides over the entire surface. A. ochraceus grows at 378C, unlike the closely related species A. westerdijkiae and A. steynii (see below). Of the other species described here, A. ochraceus most resembles A. wentii: A. ochraceus grows more rapidly on CYA and MEA than does A. wentii, but less rapidly on G25N. Conidia of A. wentii are golden yellow, not ochre. Taxonomy. Domsch et al. (1980) and Samson and Gams (1985) took up the name Aspergillus alutaceus Berk. and Curtis for this species as it predates A. ochraceus. However, this change did not meet with broad acceptance, both because of reluctance to change such a well known and apt epithet and because doubt existed as to whether these two species were synonymous. A. ochraceus was accepted in the list of Names in Current Use (Pitt and Samson, 1993). A. ochraceus is therefore the appropriate name for this species. In searches, the name A. alutaceus should also be checked in literature from about 1990 to 2000. Two closely related species, A. westerdijkiae Frisvad and Samson and A. steynii Frisvad and Samson (Frisvad et al., 2004), have recently been described (see Additional species at the end of this section). Physiology. Because Aspergillus ochraceus was confused with A. westerdijkiae and A. steynii until recently, the physiological data reported for A. ochraceus sometimes refer to these other two

318

8 Aspergillus and Related Teleomorphs

Fig. 8.17 Aspergillus ochraceus (a) colonies on CYA and MEA, 7 days, 258C; (b) heads, bar = 20 mm; (c) conidia, bar = 5 mm

species. In particular, A. ochraceus isolates grow quite strongly at 378C, indicating a maximum for growth of at least 408C. The other two species do not grow at 378C. It is not clear which species were studied when Palacios-Cabrera et al. (2005) reported no growth by A. ochraceus at either 8 or 418C on a selection of media.

It seems likely that all three species will have similar water relations, though it is unknown which species was studied in the reports that follow. Growth was reported down to 0.79 aw on glucose/fructose media and down to 0.81 aw on media based on NaCl (Pitt and Hocking, 1977). Pardo et al. (2006a) indicated germination occurred down to 0.80 aw at 20 or

8.4

Genus Aspergillus

308C and 0.85 aw at 108C. No germination occurred at 0.75 aw on barley meal extract medium (Pardo et al., 2004a) or on green coffee extract agar (Pardo et al., 2005a). The optimum aw for growth is 0.95– 0.99 aw (ICMSF, 1996a; Pardo et al., 2006a). A. ochraceus grew well between pH 3 and 10 and slowly at pH 2.2 (Wheeler et al., 1991). The decimal reduction time (D10) for Aspergillus ochraceus subjected to low dose gamma irradiation was given as 0.44 kGy on grain (O’Neill et al., 1991). Up to 4 kGy of gamma irradiation was necessary to prevent growth of A. ochraceus on barley (Szekely et al., 1991). Irradiation had no intrinsic effect on ochratoxin A production (Chelack et al., 1991). In the presence of 20% O2, Aspergillus ochraceus was able to grow in 60% but not 80% CO2 (Paster et al., 1983). Germ tube elongation was significantly inhibited by 50% CO2 at 0.90–0.95 aw and both growth and ochratoxin A production were affected by 25 and 50% CO2, regardless of the water activity (Cairns-Fuller, 2004). Like Aspergillus flavus and A. parasiticus, A. ochraceus and related species produce a bright orange reverse on AFPA, though not the same pigment (Assante et al., 1981). Growth is much slower: reading results from AFPA plates before 5 days will avoid false positives from A. ochraceus and related species (Pitt et al., 1983). A. niger has been reported to inhibit growth of A. ochraceus (Paster et al., 1992), but this does not appear to have practical applications. Mycotoxins. The toxigenic nature of Aspergillus ochraceus was discovered not as the result of a natural toxicosis but in laboratory studies on toxigenic fungi (van der Merwe et al., 1965). Three toxins were found: the major one was named ochratoxin A, while minor components of lower toxicity were named ochratoxins B and C. Subsequent work has shown that only a minority of A. ochraceus isolates are toxigenic (see Miyaki et al., 1970; Ciegler, 1972; Cvetnic, 1994; Chourasia, 1995) and that other species closely related to A. ochraceus, A. westerdijkiae and A. steynii are the main producers of ochratoxin A (Frisvad et al., 2004). It is now believed that the well-documented occurrence of ochratoxin A in temperate climate foods results from growth of Penicillium verrucosum. The properties of ochratoxin A are described under that species. Aspergillus ochraceus and related species

319

produce most of the ochratoxin A found in coffee, but it is likely that most of the ochratoxin A produced in tropical commodities is due to the growth of A. carbonarius. Possible involvement of A. ochraceus in the production of ochratoxin under tropical conditions should not be overlooked, however. Pardo et al. (2004b) found optimum ochratoxin A production by Aspergillus ochraceus on barley grains at 0.98–0.99 aw and 25–308C. Similar results were observed for maximum ochratoxin A production on irradiated green coffee beans and the highest yields occurred at 208C (Pardo et al., 2005b). The minimum water activity allowing ochratoxin A formation was reported to be 0.85 aw on sterilised coffee beans (Taniwaki et al., 1999; Pardo et al., 2005b); however, no ochratoxin A was produced at this water activity on irradiated barley grains or on grapes (Pardo et al., 2004b, c and see Pardo et al., 2006a for a recent review). The effect of pH and trace metals on the production of ochratoxin A by Aspergillus ochraceus was examined in liquid media (Mu¨hlencoert et al., 2004). At 258C, A. ochraceus synthesised ochratoxin A (up to 2 mg/l) at pH values ranging from 5.5 to 8.5, and no significant difference was observed in the amount of ochratoxin A produced at pH values within this range. Outside this pH range, however, no ochratoxin A was detected. At pH 6.5, variations in trace metal concentrations such as Zn and Fe affected ochratoxin A production (Mu¨hlencoert et al., 2004). From a study of the growth of Aspergillus ochraceus on cereals (maize and wheat) and oilseeds (peanuts, rapeseed and soybeans), it was concluded that ochratoxin production was favoured more by peanuts and soybeans than the other substrates studied (Madhyastha et al., 1990). Pardo et al. (2006b) also reported that the nature of the substrate influenced toxin production by A. ochraceus. The highest ochratoxin A yields were produced on green coffee beans (>2 mg/g), barley grains (ca. 1 mg/g), yeast extract sucrose medium (13.9 mg/g) and grape berries (ca. 3 ng/g). The toxigenic potential of the strains studied was independent of their origin (Pardo et al., 2006b). Ochratoxin A production by Aspergillus ochraceus was similar in atmospheres with 5 or 1% O2 as in air. In the presence of 20% O2, low levels of CO2 had no influence on ochratoxin A production, but it was completely inhibited by 30% CO2 in the

320

presence of 20% O2 (Paster et al., 1983). Sorbic acid had little effect on ochratoxin A production at 258C and pH 5.0 (Bullerman, 1985). The effect of heat on ochratoxin A produced in wheat by a strain of Aspergillus ochraceus was reported by Boudra et al. (1995). For dry wheat, 50% decomposition occurred in 700, 200, 12 and 6 min at 100, 150, 200 and 2508C, respectively. The addition of 50% water increased decomposition rates at 100 and 1508C, but not at higher temperatures (Boudra et al., 1995). Aspergillus ochraceus also produces penicillic acid (Frisvad et al., 2004), a mycotoxin of lesser importance described under Penicillium commune. It is produced by A. ochraceus between about 10 and 358C and down to 0.81 aw (ICMSF, 1996a). Sorbic acid (500 mg/kg) had little effect on penicillic acid production at 15–358C, but 1,000 mg/kg markedly reduced production at 15 and 258C (Gourama and Bullerman, 1988). Emodin is also produced by Aspergillus ochraceus or a sibling species and was found at low levels in chestnuts on the Canadian market (Overy et al., 2003). See A. wentii for details of emodin. Ecology. An important source of Aspergillus ochraceus and the closely related A. westerdijkiae and A. steynii is green coffee beans (Mislivec et al., 1983; Tsubouchi et al., 1985; Te´ren et al., 1997; Urbano et al., 2001; Bucheli and Taniwaki, 2002; Martins et al., 2003; Taniwaki et al., 2003; Sua´rezQuiroz et al., 2005) which may result in ochratoxin A contamination of roasted and instant coffee (Jørgensen, 2005; Clark and Snedeker, 2006). Aspergillus ochraceus and related species have also been isolated from a wide range of other foods, but are more common in dried and stored foods than elsewhere. Stored foods from which they have been isolated include soybeans (Pacin et al., 2002; Aziz et al., 2006), pepper (Gatti et al., 2003) and dried fruit (Iamanaka et al., 2005). Other foods include smoked dried fish, salted dried fish, dried beans, biltong, chickpeas, rapeseed, sesame seeds before and after fermentation, blackgram seeds and rootstock snacks (see Pitt and Hocking, 1997). Nuts are also a major source, including peanuts (Aziz et al., 2006), pecans, pistachios, hazelnuts, walnuts and betel nuts (see Pitt and Hocking, 1997). Aspergillus ochraceus and related species have been reported from cereals and cereal products, but rather infrequently. Recent records include rice (Pacin et al., 2002) and barley (Medina et al.,

8 Aspergillus and Related Teleomorphs

2006), though levels in Europe are lower than for some other fungal species (Sepitkova and Jesenska, 1986), maize (Sepulveda and Piontelli, 2005; Magnoli et al., 2006b, 2007a for a recent review) and maize meal (Souza et al., 2003), corn snacks (Adebajo et al., 1994), wheat (Aziz et al., 2006; Hajjaji et al., 2006), flour (Halt et al., 2004) and bran (Dragoni et al., 1979). Aspergillus ochraceus and related species were the major contaminant in Bhutanese cheese (Sinha and Ranjan, 1991). They have also been reported from cheese in the temperate zone (see Pitt and Hocking, 1997). Electron beam irradiation of 0.42 kGy enabled a greatly increased shelf life for refrigerated Cheddar cheese surface inoculated with A. ochraceus (Blank et al., 1992). Other less common sources include spices (Almela et al., 2007), cassava (Souza et al., 2003), black olives and processed meats (see Pitt and Hocking, 1997). We isolated Aspergillus ochraceus and related species from a variety of Southeast Asian commodities, including maize, peanuts, soybeans and other beans, cashews and sorghum. Its presence or absence in any sample probably related to length of storage rather than geographical location or other factors (Pitt et al., 1993, 1994, and 1998a). Rots in garlic have been reported to be caused by Aspergillus ochraceus, but the species responsible is A. alliaceus Thom and Church (Snowdon, 1991). Additional species. Aspergillus westerdijkiae Frisvad and Samson (Frisvad et al., 2004) is closely related to A. ochraceus with perhaps slightly faster growth rates on CYA at 258C. No growth occurs at 378C and sclerotia, which are sparsely produced, remain white to cream with age. Conidia of A. westerdijkiae are consistently finely roughened, compared with smooth conidia in A. ochraceus. A. westerdijkiae produces large amounts of ochratoxin A. It also produces penicillic acid, xanthomegnin, viomellein and vioxanthin. Some physiological studies conducted on isolates identified as A. ochraceus undoubtedly relate to A. westerdijkiae (Frisvad et al., 2004). Because of its ability to synthesise high levels of ochratoxin A, A. westerdijkiae is of importance in commodities such as coffee, cereals and beverages. Additional species. Aspergillus steynii Frisvad and Samson (Frisvad et al., 2004) is also closely related to A. ochraceus. The absence of growth at 378C, ellipsoidal conidia and pale yellow conidia distinguish this species from A. ochraceus. Like A.

8.4

Genus Aspergillus

westerdijkiae, A. steynii produces copious amounts of ochratoxin A, along with penicillic acid and xanthomegnins. Its broad distribution in commodities such as coffee, soybeans and rice makes A. steynii an important species (Frisvad et al., 2004). References. Raper and Fennell (1965); Domsch et al. (1980), under Aspergillus alutaceus; Christensen (1982); Klich (2002) (all under A. ochraceus); Frisvad et al. (2004) (for A. ochraceus, A. westerdijkiae and A. steynii).

321

Aspergillus parasiticus Speare

Fig. 8.18

Colonies on CYA 50–70 mm diam, plane, low, dense and velutinous; mycelium inconspicuous, white; conidial heads in a uniform, dense layer, dark yellowish green (29-30D-F6-8); sclerotia occasionally produced; reverse uncoloured or brown. Colonies on MEA 50–65 mm diam, generally similar to those on CYA but usually less dense and with reverse uncoloured. Colonies on G25N

Fig. 8.18 Aspergillus parasiticus (a) colonies on CYA and MEA, 7 days, 258C; (b, c) heads, bars ¼ 10 mm; (d) conidia, bar ¼ 5 mm

322

20–40 mm diam, plane, low and velutinous, generally similar to those on CYA; reverse uncoloured, yellow or brown. No growth at 58C. At 378C, colonies covering the available area, similar to those on CYA at 258C, or with conidia deeper green or brownish; reverse pale. Sclerotia occasionally produced, white at first, becoming black, spherical, 400–800 mm diam. Teleomorph not known. Conidiophores borne from subsurface or surface hyphae, stipes 250–500 mm long, with colourless or pale brown, smooth walls; vesicles spherical, 20–35 mm diam, fertile over three quarters of the surface, mostly bearing phialides only, but in some isolates up to 20% of heads bearing metulae as well; phialides 7–11 mm long; conidia spherical, mostly 4.0–6.0 mm diam, with distinctly roughened walls, usually borne in radiate heads. Distinctive features. Aspergillus parasiticus shares with A. flavus fast growth at both 25 and 378C and distinctive bright yellow green conidial colours. A. parasiticus differs from A. flavus by the production of spherical conidia with heavy, rough walls, while A. flavus conidia are more variable in shape and have relatively thin, usually finely roughened walls. Taxonomy. Production of phialides only was the principal character used by Raper and Fennell (1965) to distinguish Aspergillus parasiticus from A. flavus. Klich and Pitt (1985, 1988) showed that this criterion was unsatisfactory, as some A. flavus isolates produce very few metulae. Careful study showed several differences between the two species, the most useful of which are mentioned above. The criteria used here provide a 100% correlation with mycotoxin production (see below). Kurtzman et al. (1986b) reduced Aspergillus parasiticus to the status of subspecies, as A. flavus subspecies parasiticus (Speare) Kurtzman et al., but as discussed under Aspergillus flavus, compelling reasons exist for maintaining distinct species names for these taxa, as has been done here. A. sojae, reduced to the status of variety by Kurtzman et al. (1986b), appears to be a domesticated form of A. parasiticus. As with A. oryzae, A. sojae is widely used in food fermentations, and recognition of it as a separate species is a practical necessity. Random amplification of polymeric DNA (RAPD) has been used to distinguish between A. parasiticus and A. sojae (Yuan et al., 1995).

8 Aspergillus and Related Teleomorphs

Physiology. A number of earlier studies reported to be on the physiology of Aspergillus flavus have used misidentified strains and have really dealt with A. parasiticus (Pitt, 1993b). These two species are physiologically very similar, so that the studies reported under A. flavus are probably equally applicable to A. parasiticus. Growth of A. parasiticus has been reported over the range 12–428C, with an optimum at 328C (ICMSF, 1996a). A minimum aw for growth of 0.82 at 258C, 0.81 at 308C and 0.80 at 378C, similar to A. flavus, has been reported (Pitt and Miscamble, 1995). Data from that paper were used to provide a predictive model for A. parasiticus growth in relation to aw and temperature (Gibson et al., 1994). Growth of A. parasiticus occurred over the pH range 2.4–10.5 at 25, 30 and 378C, but growth did not occur at pH 2.2 at 258C. Optimal growth occurred over a broad range from pH 3.5–8, with growth at pH 10.5 still more than half that under optimal conditions (Wheeler et al., 1991). High levels of sorbic acid (1,000 mg/kg) were needed to inhibit growth of A. parasiticus on synthetic media at pH 5 (Bullerman, 1983). Conidia of Aspergillus parasiticus have a low heat resistance: a D55 up to 9 min at pH 7 in phosphate buffer (Doyle and Marth, 1975a). However, values were much higher if aw was reduced: a D55 of 230 min in 16% NaCl (0.90 aw), 210 min in 60% glucose (0.85 aw) and 200 min in 60% sucrose (0.90 aw) (Doyle and Marth, 1975b; ICMSF, 1996a). D values for the effect of gamma irradiation on conidia of A. parasiticus ranged from 0.3 to 0.6 kGy (ICMSF, 1996a). Like Aspergillus flavus, A. parasiticus colonies exhibit a brilliant orange yellow reverse colouration after 42–48 h incubation on AFPA (Pitt et al., 1983). Mycotoxins. The important differences in mycotoxin production between Aspergillus parasiticus and A. flavus are that A. parasiticus produces G as well as B aflatoxins, that A. parasiticus isolates often produce aflatoxins in much higher concentrations (Pitt, 1993b) and that nontoxigenic A. parasiticus strains are rare (Horn et al., 1994; Tran-Dinh et al., 1999). A. parasiticus does not produce cyclopiazonic acid (Vaamonde et al., 2003; Horn, 2003; Frisvad et al., 2005a) but does produce kojic acid, a metabolite of low toxicity now finding wide use in medicine and cosmetics (El-Aasar, 2006).

8.4

Genus Aspergillus

The conditions under which Aspergillus parasiticus produces aflatoxins are similar to those reported for A. flavus. The best available figures indicate production over the temperature range 12–408C, down to 0.86 aw, and over the pH range 3–8 or higher (ICMSF, 1996a). The ratio of aflatoxin G1 production relative to B1 on a rice substrate was higher at lower temperatures (at 15– 188C, G1:B1 was 1.3–1.5) and much reduced at high temperatures (288C, 0.24 and 328C, 0.06) (Sorenson et al., 1966; ICMSF, 1996a). The G1:B1 ratio was up to 2.5 at 158C on peanuts (Diener and Davis, 1967). In the presence of 20% O2, at least 60% CO2 was needed to prevent production of aflatoxins on a moist peanut substrate at 308C, though the production in 60% CO2 was only 10% of that in air (Landers et al., 1967). In reduced levels of O2 in N2, 5% O2 caused aflatoxin production to be reduced to 30% of that in air, while in 1% O2 production was only 1% of that in air (Landers et al., 1967). Other aspects of aflatoxins, including toxicity and control, are discussed under A. flavus. Ecology. Records of Aspergillus parasiticus from foods are relatively rare. Although A. parasiticus is certainly widely distributed in soils and foodstuffs in the United States, Latin America, South Africa, India and Australia, it is essentially unknown in Southeast Asia (Pitt et al., 1993, 1994, and 1998a). Like Aspergillus flavus, A. parasiticus is a tropical and subtropical species, less prevalent in warm temperate zones and rare in the cool temperate regions of the world. Its absence from Southeast Asia indicates a limited distribution. In soils, populations of A. parasiticus are generally associated with peanut cultivation in the United States and Australia (Carter et al., 1998; Horn, 2003). This species has only been reported in low densities in corn fields (Horn, 2003). The most important food source is peanuts, in which we believe A. parasiticus is endemic, but about which reliable information is surprisingly limited. Klich and Pitt (1988) reported studying isolates from peanuts from the United States, Australia and Uganda. This species has been reported in peanuts in Argentina (Vaamonde et al., 2003) and Botswana (Mphande et al., 2004). Isolations of A. parasiticus from Australian peanuts are now more numerous than those of A. flavus (our data). Other types of nuts may also be infected: hazelnuts and walnuts, pistachios and pecans (see Pitt and Hocking, 1997).

323

Maize is perhaps a source (Aja-Nwachukwu and Emejuaiwe, 1994), but we believe that Aspergillus parasiticus is much less common on this commodity than A. flavus. A recent survey of Italian maize revealed that most contamination was due to A. flavus and A. parasiticus occurred in low frequency (Giorni et al., 2007). A. parasiticus has been reported at low frequency in rice and other rice derivatives (Sales and Yoshizawa, 2005b). Other reported sources include soybeans (Vaamonde et al., 2003), processed meats (El-Tabiy, 2006), black pepper (Gatti et al., 2003; Mandeel, 2005) as well as rootstock snacks, herbal drugs, amaranth seeds and pearl millet (see Pitt and Hocking, 1997). References. Raper and Fennell (1965); Domsch et al. (1980); Christensen (1981); Klich and Pitt (1988); Klich (2002).

Aspergillus penicillioides Spegazzini

Fig. 8.19

Colonies on CYA up to 5 mm diam, sometimes only microcolonies, of white mycelium only. Growth on MEA usually limited to microcolonies, occasionally colonies up to 5 mm diam, similar to those on CYA. Colonies on G25N 8–15 mm diam, plane or centrally raised, sometimes sulcate or irregularly wrinkled, texture velutinous or lightly floccose; mycelium usually inconspicuous, white; conidial production moderate, coloured dull green to dark green; reverse pale to dark green. No growth at 5 or 378C on CYA. Colonies on CY20S varying from microcolonies up to 10 mm diam, similar to those on CYA, occasionally some dull green conidial production but conidiophores poorly formed; reverse pale. Colonies on MY50G 10– 16 mm diam, plane or umbonate, relatively sparse, velutinous to floccose; conidial production moderate, greyish green to dull green; reverse pale. Conidiophores borne from surface or aerial hyphae, showing optimal development on G25N, stipes (150–)300–500 mm long, sometimes sinuous, with colourless, thin, smooth walls, enlarging gradually from the base, then rather abruptly to pyriform or spathulate vesicles; vesicles mostly 10–20 mm diam, usually fertile over two thirds of the area, bearing phialides only; phialides 8–11 mm long; conidia borne as ellipsoids, at maturity ellipsoidal to subspheroidal, 4.0–5.0 mm diam, with spinose walls, typically borne in radiate heads, uncommonly in loose columns also.

324

8 Aspergillus and Related Teleomorphs

Fig. 8.19 Aspergillus penicillioides (a) colonies on CYA and MY50G, 7 days, 258C; (b, c) heads, bars = 10 mm; (d) conidia, bar = 5 mm

Distinctive features. In common with Aspergillus restrictus, A. penicillioides grows very slowly under all standard conditions and produces green conidia. It differs from A. restrictus by very weak growth on CYA and MEA, by forming radiate conidial heads from spathulate vesicles, fertile over more than the upper half; and by bearing conidia as ellipsoids, which usually separate in liquid mounts.

Physiology. As is clear from the description above, this species grows very poorly at high aw. The optimal aw for growth is 0.89 at 308C in glucose/fructose media of pH 5.5 (Gock et al., 2003) or 0.91–0.93 at 258C on media containing either glucose/fructose or NaCl, at pH 6.5 (Andrews and Pitt, 1987). At 258C, Aspergillus penicillioides is capable of germination but not growth at 0.70 aw and pH

8.4

Genus Aspergillus

4.5–5.5 in media containing glucose/fructose (Gock et al., 2003). No germination is observed at this aw at pH values above 5.5 at 25, 30 or 378C. On NaClbased media, the limit is 0.75 aw (Andrews and Pitt, 1987). The minimum at 20, 30 and 348C is higher, 0.76–0.79 aw on glucose/fructose and 0.80–0.84 on NaCl. At 378C, minimum figures are 0.79 and 0.88 aw, respectively (Wheeler et al., 1988b). In natural substrates, A. penicillioides is capable of growth at much lower water activities: we have reproducible evidence of its ability to multiply at 0.68 aw in wheat and pet food within 6 months (our unpublished observations). Growth temperatures have not been accurately reported: from the above, growth does not occur at 378C on CYA, but occurs readily at this temperature at lower aw (Wheeler et al., 1988b; Gock et al., 2003). Growth occurs down to 158C or lower, at 0.95–0.90 aw at least (Wheeler et al., 1988b). Mycotoxins. This species has not been reported to produce mycotoxins. Ecology. Reports of Aspergillus penicillioides in foods are quite rare, primarily because it does not grow on the media commonly used for fungal isolation and enumeration. At best, development on DRBC or other high water activity isolation media is poor. Greatly improved results can be obtained if a low aw medium such as DG18 is used (Hocking, 1981). It is also true that this species has frequently been misidentified as A. restrictus. For example, A. restrictus was reported to be a major cause of loss of germinability in wheat stored just above the safe moisture content (Tuite and Christensen, 1957; Christensen, 1963). In fact the species responsible is A. penicillioides. In this laboratory, we have isolated Aspergillus penicillioides, often in very high numbers, from a wide variety of foods, including flour, dried fruit and dried fish, and from spices, including pepper and dried chillies. Other reported isolations have come from milled rice (Taligoola et al., 2004), fermented and cured meats, cocoa and peanuts (see Pitt and Hocking, 1997) and salted dried fish (Nketsia-Tabiri et al., 2003; Gra¨u et al., 2007). This species was quite competitive at 308C on glucose media, but was less so on NaCl media or salt fish, or at other temperatures (Wheeler and Hocking, 1993). From Southeast Asian commodities, we isolated Aspergillus penicillioides quite frequently. Although

325

the proportion of samples infected was low, individual samples often showed high infection rates, no doubt reflecting prolonged storage at water activities just above safe limits. A. penicillioides was commonly isolated from soybeans in Thailand and Indonesia, where 6 and 17% of samples were contaminated, respectively. Black pepper from the Philippines showed up to 60% infection in contaminated samples and 2% infection in peppercorns overall. Lower levels of infection were found in paddy and milled rice, various other kinds of beans including mung beans and peanuts (Pitt et al., 1993, 1994, 1998a and our unpublished data). References. Raper and Fennell (1965); Pitt and Samson (1990c); Klich (2002).

Aspergillus restrictus G. Sm.

Fig. 8.20

Colonies on CYA 6–12 mm diam, sulcate or wrinkled, low, dense and velutinous; mycelium inconspicuous, white; conidial heads often poorly formed, sparse to numerous, in the latter case dull green; reverse pale to very dark green. Colonies on MEA 6–12 mm diam, occasionally smaller, similar to those on CYA or centrally raised, conidial production heavy but heads poorly formed, coloured dull green to dark green; reverse usually pale. Colonies on G25N 10–15 mm diam, plane or umbonate, usually similar to those on MEA, but heads well formed; reverse sometimes dark green. No growth at 5 or 378C on CYA . Colonies on CY20S 16–20 mm diam, generally similar to those on G25N apart from slightly more rapid growth. Colonies on MY50G 12–16 mm diam, plane or umbonate, with aerial growth and conidial production usually sparse, coloured greyish green to dull green; reverse pale. Conidiophores borne from surface hyphae, developing optimally on CY20S, stipes 75–200 mm long, sometimes sinuous, with colourless, thin, smooth walls, enlarging from the base gradually then abruptly to pyriform vesicles; vesicles 10–18 mm diam, fertile over the upper hemisphere or less, bearing phialides only; phialides crowded, 8–10 mm long; conidia borne as cylinders, in long, appressed columns, adhering in liquid mounts, when mature nearly cylindrical to doliiform (barrel shaped), 4.0–6.0 mm long, with rough walls, borne in long columns under favourable growth conditions.

326

8 Aspergillus and Related Teleomorphs

Fig. 8.20 Aspergillus restrictus (a) colonies on CYA and MEA, 7 days, 258C; (b, c) heads, bars = 10 mm; (d) conidia, bar = 5 mm

Distinctive features. In common with Aspergillus penicillioides, A. restrictus grows very slowly under all conditions and produces green conidia. It differs from A. penicillioides by vesicles which are fertile over the upper half or less and by conidia borne as cylinders and adhering in long columns, usually persisting in liquid mounts. A. restrictus grows more rapidly than A. penicillioides on CY20S. Physiology. Smith and Hill (1982) reported that the temperature range for growth of Aspergillus restrictus was minimum 98C, optimum 308C and

maximum 408C. Growth of this species has been observed down to 0.75 aw (Snow, 1949; Pelhate, 1968). This species grows strongly in 1% O2 but not at all in 0.1% O2 (Hocking, 1990). Mycotoxins. This species has not been reported to produce mycotoxins. Ecology. Considering the slow growth rate of this species, and its inconspicuous habit, Aspergillus restrictus has been isolated from foods quite frequently. Most records have come from dried foods: wheat, rice, maize, dried beans, pecans,

8.4

Genus Aspergillus

peppercorns, salted dried fish, cured meats, Japanese traditional noodles, spices, pepper, carriers for dough during breadmaking, dried prunes and health foods (see Pitt and Hocking, 1997). A. restrictus has been reported recently in cashew nuts (Adebajo and Diyaolu, 2003), coffee beans (Ahmad and Magan, 2003), cocoa beans (Wojcik-Stopczynska, 2006) and cured fish (Nketsia-Tabiri et al., 2003). We isolated Aspergillus restrictus quite frequently from Southeast Asian commodities. As with A. penicillioides, the proportion of samples infected was often low, but with individual samples showing high infection rates, reflecting prolonged storage. The highest levels of A. restrictus found were in samples of red rice from Indonesia. Soybeans from the Philippines, Indonesia and Thailand showed low levels of infection (see Pitt and Hocking, 1997). Mung beans, red beans and cowpeas showed up to 24% infection in individual samples. Black pepper from the Philippines showed up to 100% infection in contaminated samples. Only 1 sample of kemiri nuts (of 20) from Indonesia was infected, but this species infected 30% of nuts in that sample. Low levels of infection occurred in paddy and milled rice, maize and peanuts (Pitt et al., 1993, 1994, 1998a and our unpublished data). References. Raper and Fennell (1965); Domsch et al. (1980); Pitt and Samson (1990c); Klich (2002).

Aspergillus sydowii (Bainier & Sartory) Thom & Church Fig. 8.21 Colonies on CYA 18–30 mm diam, plane or lightly sulcate, low to moderately deep, dense and velutinous to somewhat floccose; mycelium white; conidial heads sparse to quite dense, dark turquoise to dark green, especially in marginal areas, centrally sometimes buff to orange brown; dark brown exudate and/or soluble pigment sometimes produced; reverse pale to orange brown. Colonies on MEA 16–25 mm diam, plane, dense, velutinous to lightly floccose; mycelium inconspicuous, white; conidial heads numerous, coloured like those on CYA; reverse pale. Colonies on G25N 15–20 mm diam, plane, dense; mycelium white; often heavily sporing, dull green, blue or brown; reverse pale or yellowish. Usually no growth at 58C, occasionally germination. At 378C, no growth or colonies up to 10 mm diam formed .

327

Conidiophores borne from surface or aerial hyphae, stipes 250–500 mm long, often sinuous, with heavy, pale brown, smooth walls; vesicles only slightly swollen, club shaped, 10–20 mm diam, bearing metulae and phialides over the upper two thirds to three quarters; smaller conidiophores also produced from aerial hyphae, ranging down to tiny monoverticillate penicilli; metulae 4–7 mm long; phialides 7–10 mm long; conidia spherical, 2.5–3.5 mm diam, with spinose walls, from the larger conidiophores borne in radiate heads. Distinctive features. Aspergillus sydowii grows slowly at 258C and often not at all at 378C, produces heads with both metulae and phialides and blue conidia. Vesicles on the larger stipes are small and club shaped, and diminutive penicilli are also formed. Taxonomy. The taxonomy of Aspergillus versicolor and related species, including A. sydowii, was reviewed and revised by Klich (1993). Physiology. Closely related to Aspergillus versicolor, A. sydowii can be expected to have similar physiological properties. Snow (1949) reported 0.78 aw to be the minimum for growth on a natural substrate; Wheeler and Hocking (1988) gave the same figure for growth on a glucose/fructose medium and 0.81 for growth on NaCl-based media. Only 0.7% of A. sydowii conidia survived heating at 508C for 10 min (Pitt and Christian, 1970). Mycotoxins. No significant mycotoxins have been reported from this species. Ecology. Widely distributed, Aspergillus sydowii is a storage fungus. Most isolations have come from dried foods, including various nuts: peanuts, pistachios, coconut, hazelnuts, walnuts and pecans (see Pitt and Hocking, 1997); fermented milk (Ahmed and Adbel-Sater, 2003), coffee beans (Silva et al., 2003), dried fish, paddy rice, stored rice, soybeans, cured meats and biltong, dried beans, health foods, red pepper powder (see Pitt and Hocking, 1997) and spices (Ramaswamy et al., 2004). It appears to be relatively uncommon in cereals, but has been isolated from barley, wheat and flour (see Pitt and Hocking, 1997), maize and soy beans (Sepulveda and Piontelli, 2005). It has been reported as one cause of spoilage in fresh breadfruit (Omobuwajo and Wilcox, 1989). Aspergillus sydowii was quite common in Southeast Asian commodities. We isolated it from 80% of pepper samples from Indonesia and 31% of samples from the Philippines. Levels in Indonesian

328

8 Aspergillus and Related Teleomorphs

Fig. 8.21 Aspergillus sydowii (a) colonies on CYA and MEA, 7 days, 258C; (b, c) heads, bars = 10 mm; (d) conidia, bar = 5 mm

coriander were also high. It was significant (1–2% overall) in cashews and copra from Thailand; soybeans from the Philippines; and kemiri nuts, sorghum and black soybeans from Indonesia. It was also present at low levels in maize, peanuts, mung beans, paddy rice and milled rice from one country or more (Pitt et al., 1993, 1994, 1998a and our unpublished data). References. Raper and Fennell (1965); Domsch et al. (1980); Klich (1993, 2002).

Aspergillus tamarii Kita

Fig. 8.22

Colonies on CYA 50–65 mm diam, plane, velutinous to lightly floccose; mycelium inconspicuous, white; conidial heads abundant, olive brown; reverse uncoloured. Colonies on MEA 55–65 mm diam, similar to those on CYA but relatively sparse and with olive conidia; reverse uncoloured. Colonies on G25N 35–40 mm diam, similar to those on CYA but coloured deep olive brown; reverse

8.4

Genus Aspergillus

329

Fig. 8.22 Aspergillus tamarii (a) colonies on CYA and MEA, 7 days, 258C; (b, c) heads, bars = 20 mm; (d) conidia, bar = 5 mm

uncoloured. No growth at 58C. At 378C, colonies 50–65 mm diam, much more dense than on CYA at 258C, low and velutinous, with conidia coloured coffee brown; reverse pale. Conidiophores borne from subsurface or surface hyphae, stipes 300–1,500 mm long, colourless, usually with rough, thin walls; vesicles spherical to subspheroidal, 25–50 mm diam, fertile virtually all over, bearing both metulae and phialides or less commonly phialides alone; metulae 12–20 mm long; phialides 9–12 mm long; conidia spherical to subspheroidal,

5–8 mm diam, brown, with characteristic thick, rough to spiny walls, borne in radiate heads. Distinctive features. Aspergillus tamarii shows an unmistakable resemblance to A. flavus and A. parasiticus. The main distinguishing features are that conidia of A. tamarii are coloured olive to brown on CYA and MEA and are larger, with thick, conspicuously roughened walls. On AFPA, A. tamarii produces a deep brown reverse colouration, in contrast to the orange yellow of A. flavus and A. parasiticus. This is a useful diagnostic aid.

330

Taxonomy. Domsch et al. (1980) revived the name Aspergillus erythrocephalus Berk. and Curtis for this species, but other authorities, e.g. Christensen (1981), maintained the use of A. tamarii. Examination of the type specimen of A. erythrocephalus (K.A. Seifert and R.A. Samson, unpublished) showed that it is distinct from A. tamarii. In the List of Names in Current Use (Pitt and Samson, 1993), A. tamarii is maintained as the correct name for this species. Physiology. Less has been published on the physiology of Aspergillus tamarii than on A. flavus, but it appears from growth data that gross physiological behaviour of the two species, such as temperature relations, will be similar. Ayerst (1969) reported that A. tamarii was capable of growth down to 0.78 aw at 338C, a figure identical with that which he obtained for A. flavus. A. tamarii can degrade caffeine in coffee residues (Gutie´rrez-Sa´nchez et al., 2003). Mycotoxins. Aspergillus tamarii does not produce aflatoxins, but does produce cyclopiazonic acid (Dorner, 1983). This toxin is discussed in more detail under P. camemberti. A. tamarii may have been the cause of ‘‘kodua poisoning’’ from kodo millet seed (Paspalum scrobiculatum) in India (Rao and Husain, 1985). The alkaloid fumiclavine A was also found in kodo millet seed from that outbreak, and it may have contributed to the toxicity (Janardhanan et al., 1984). A. tamarii was identified as the source of both toxins. Antony et al. (2003) demonstrated that exposure to cyclopiazonic acid through ingestion of contaminated kodo millet may cause acute hepatotoxicity in men and animals. A. tamarii also produces kojic acid, a compound of low toxicity (Manabe et al., 1984). Ecology. While not as universally encountered in foods as Aspergillus flavus, A. tamarii is nevertheless of widespread occurrence in tropical and subtropical regions. Like A. flavus, it occurs commonly in nuts and oilseeds (see Pitt and Hocking, 1997). Isolations from cereals have been infrequent. A. tamarii has been recorded in wheat, barley and sorghum (see Pitt and Hocking, 1997). Other sources include soybeans and maize (Amusa et al., 2005, Sepulveda and Piontelli, 2005), green coffee beans (Silva et al., 2003; Tharappan et al., 2006), meat products (Mohamed and Hussein, 2004), spices, peppercorns, salted, smoked and dried fish and a variety of tropical products including cocoa, palm kernels, maize, yams and melon ball snacks (see Pitt and Hocking, 1997).

8 Aspergillus and Related Teleomorphs

Adeniji (1970a) reported isolation of Aspergillus tamarii from rotting yams, and it was reported as one cause of kernel rot in cashews (Esuruoso, 1974). Amusa et al. (2003) isolated this species from African star apple. Southeast Asian food commodities we examined were commonly infected with Aspergillus tamarii. Indonesian pepper was severely contaminated, although peppercorns harboured slightly lower rates of infection. Levels in peanuts were high, but lower in maize. A. tamarii was also found on kemiri nuts from Indonesia, peppercorns and soybeans from the Philippines, copra and black beans from Thailand and paddy rice from Indonesia. Other commodities, including cashews, mung beans and soybeans from Thailand and Indonesia, mung beans and paddy and milled rice from the Philippines and cowpeas from Indonesia showed low levels of infection (Pitt et al., 1993, 1994, 1998a and our unpublished data). References. Raper and Fennell (1965); Domsch et al. (1980), under Aspergillus erythrocephalus; Christensen (1981); Klich (2002).

Aspergillus terreus Thom

Fig. 8.23

Colonies on CYA 40–50 mm diam, plane, low and velutinous, usually quite dense; mycelium white; conidial production heavy, pale pinkish brown to blonde; reverse pale to dull brown or yellow brown. Colonies on MEA 40–60 mm diam, similar to those on CYA or less dense. Colonies on G25N 18–22 mm diam, plane or irregularly wrinkled, low and sparse; conidial production light, pale brown; brown soluble pigment sometimes produced; reverse brown. No growth at 58C. Colonies at 378C growing very rapidly, 50 mm or more diam, of similar appearance to those on CYA at 258C. Conidiophores borne from surface hyphae, stipes 100–250 mm long, smooth walled; vesicles 15–20 mm diam, fertile over the upper hemisphere, with densely packed, short, narrow metulae and phialides, both 5–8 mm long; conidia spherical, very small, 1.8–2.5 mm diam, smooth walled, at maturity borne in long, well defined columns. Distinctive features. Velutinous colonies formed at both 25 and 378C, uniformly brown, with no other colouration, and minute conidia borne in long columns make Aspergillus terreus a distinctive species.

8.4

Genus Aspergillus

331

Fig. 8.23 Aspergillus terreus (a) colonies on CYA and MEA, 7 days, 258C; (b, c) heads, bars = 10 mm; (d) conidia, bar = 5 mm

Physiology. Growth data reported here indicate that Aspergillus terreus is thermophilic, but there appear to be no published data on this aspect of its physiology. A minimum aw for growth of 0.78 at 378C has been reported (Ayerst, 1969). Mycotoxins. Aspergillus terreus produces a wide range of metabolites (Frisvad and Samson, 1991; Samson and Frisvad, 2004), but of these only territrems appear to have significant toxicity. Territrems

are tremorgenic toxins which, unlike all other tremorgens, lack a moiety containing nitrogen. Maximum toxin production occurred at 288C following 12 days’ incubation in potato dextrose liquid medium (Fang and Peng, 2003). Territrem B synthesis increased at alkaline pH. Immunoelectron microscopy revealed that the toxin was localised in the cytoplasm of conidia after a minimum of 4 days growth on potato dextrose medium. No toxin was

332

detected in the fungal hyphae (Peng et al., 2004). These toxins have not been implicated in disease (Cole and Dorner, 1986; Ling, 1994) and their importance is a matter for conjecture. Ecology. Although not usually a cause of spoilage in foods, Aspergillus terreus is of common occurrence. It is well adapted to growth in stored foods in tropical and subtropical climates. It commonly occurs in nuts, e.g. pistachios (Heidarian et al., 2006), peanuts, hazelnuts, walnuts and pecans (see Pitt and Hocking, 1997). Reports from cereals include maize (Bhattacharya and Raha, 2002; Sepulveda and Piontelli, 2005) and maize meal (Souza et al., 2003), wheat (Hajjaji et al., 2006), barley, paddy rice and parboiled rice (see Pitt and Hocking, 1997). A. terreus has also been reported from animal feeds containing a mixture of cereals (Magnoli et al., 2005) and silage (El-Shanawany et al., 2005). This species has been found in cassava flour (Souza et al., 2003), flour and refrigerated dough products, pasta, miso and soy sauce (see Pitt and Hocking, 1997). Other records include soybeans (Sepulveda and Piontelli, 2005), chickpeas and mungbeans (Javaid et al., 2005), rapeseed, blackgram seeds, dried beans and peas, meat and meat products, biltong (see Pitt and Hocking, 1997), spices (Mandeel, 2005), stored coffee beans (Ahmad and Magan, 2003), dry salted fish (Gra¨u et al., 2007), cheese and curds (Kumaresan et al., 2003). Pathogenicity. Aspergillus terreus is responsible for about 10% of cases of human pulmonary aspergillosis, and it is regularly implicated in a wide range of other conditions including cutaneous and ophthalmic infections. German Shepherd dogs appear to be particularly susceptible to A. terreus infections (De Hoog et al., 2000). References. Raper and Fennell (1965); Domsch et al., (1980); Klich (2002). Aspergillus ustus (Bainier) Thom & Church Fig. 8.24 Colonies on CYA 30–40 mm diam, plane or lightly sulcate, dense, sometimes with a floccose overlay; mycelium white to greyish; conidial production sparse to quite heavy, pure grey to brownish grey; bright yellow soluble pigment usually produced; reverse greyish brown and often dull to bright yellow from soluble pigments as well.

8 Aspergillus and Related Teleomorphs

Colonies on MEA 40–50 mm diam, low, plane, dense and velutinous, or lightly floccose; mycelium white; conidial production moderate, olive brown or greyer; reverse pale green or dull brown. Colonies on G25N 10–14 mm diam, low and dense; reverse greenish or brown. At 58C, sometimes germination by a proportion of conidia. At 378C, colonies usually 30–50 mm diam, similar to those on CYA at 258C, or with brown conidia; reverse brown. Conidiophores borne from surface or aerial hyphae, stipes 100–300 mm long, sometimes curved or sinuous, with brown walls; vesicles spherical to pyriform, 10–16 mm diam, fertile over the upper two-thirds, bearing metulae and phialides, both 5–7 mm long; conidia spherical, 3.5–4.5 mm diam, brown, with rough to very rough walls, borne in small, densely packed radiate heads. Distinctive features. Aspergillus ustus is distinguished by grey conidia, small heads and vesicles bearing metulae and phialides. Growth rates at 25 and 378C on CYA are similar. Physiology. From the growth data above, Aspergillus ustus grows well at high temperatures and, interestingly, is one of the very few Aspergillus species we have encountered which can grow at low temperatures. A. ustus was isolated from deep sea (5,000 m) sediments in the Central Indian Basin. Although this species grew at 5 and 308C it was only able to synthesise an alkaline, cold tolerant protease at 308C and 1 bar pressure (Damare et al., 2006). It is not xerophilic. Mycotoxins. This species produces a number of secondary metabolites (Frisvad and Samson, 1991), but has not been reliably reported to produce significant mycotoxins. Ecology. Although Aspergillus ustus has been reported from a wide range of foods, occurrence is rather infrequent. Recent records include soybeans (Tariq et al., 2005), cashews (Freire and Kozakiewicz, 2005), almonds (Giridhar and Reddy, 2001) and grapes (Bau et al., 2006). A. ustus has also been reported in wheat, barley, flour, peanuts, walnuts, pecans, betel nuts, sago, frozen meat, biltong and cheese (see Pitt and Hocking, 1997). It has not been reported to cause food spoilage. References. Raper and Fennell (1965); Domsch et al. (1980); Klich (2002).

8.4

Genus Aspergillus

333

Fig. 8.24 Aspergillus ustus (a) colonies on CYA and MEA, 7 days, 258C; (b, c, d) heads, bars = 10 mm; (e) conidia, bar = 5 mm

Aspergillus versicolor (Vuill.) Tirab.

Fig. 8.25

Colonies on CYA 16–25 mm diam, plane or lightly sulcate, low to moderately deep, dense; mycelium white to buff or orange; conidial heads sparse to quite densely packed, greyish green; pink to wine red exudate sometimes produced; reverse brownish orange or reddish brown. Colonies on MEA 12– 25 mm diam, low, plane, and dense, usually velutinous; mycelium white to buff; conidial heads numerous, radiate, dull green; reverse yellow brown to orange brown. Colonies on G25N 10–18 mm diam, plane or umbonate, dense, of white, buff or yellow mycelium; reverse pale, yellow

brown or orange brown. No growth at 58C. Usually no growth at 378C, occasionally colonies up to 10 mm diam formed. Conidiophores borne from surface or aerial hyphae, stipes 300–600 mm long, with heavy yellow walls; vesicles variable, the largest nearly spherical, 12–16 mm diam, fertile over the upper half to two-thirds, the smallest scarcely swollen at all and fertile only at the tips, bearing closely packed metulae and phialides, both 5–8 mm long; conidia mostly spherical, very small, 2.0–2.5 mm diam, with walls finely to distinctly roughened or spinose, borne in radiate heads. Distinctive features. Aspergillus versicolor grows slowly, produces both metulae and phialides from

334

8 Aspergillus and Related Teleomorphs

Fig. 8.25 Aspergillus versicolor (a) colonies on CYA and MEA, 7 days, 258C; (b, c, d) heads, bars = 10 mm; (e) conidia, bar = 5 mm

small vesicles and green conidia. Growth at 378C is weak or absent. This species is remarkable for the wide range of mycelial and reverse pigmentation it may produce, especially if cultures are incubated for 14 days or so. Taxonomy. Klich (1993) reviewed and revised the taxonomy of Aspergillus versicolor and related species. Physiology. Aspergillus versicolor is a mesophile. Its minimum temperature for growth was reported

to be 98C at 0.97 aw, its maximum 398C at 0.87 aw and optimum 278C at 0.98 aw (Smith and Hill, 1982). Fomicheva et al. (2006) reported minimum growth on CYA at 58C, maximum growth between 35 and 378C and optimal growth between 25 and 278C. The minimum aw for growth was reported to be 0.78 (Snow, 1949) or 0.80 (Pitt and Christian, 1968), both at 258C. Growth rates at various water activities were little affected by the use of NaCl or

8.4

Genus Aspergillus

glycerol as controlling solute, or adjusting pH to 4.0 or 6.5 (Avari and Allsopp, 1983). Addition of NaCl to CYA to a concentration of 3.5% (0.98 aw) favoured the growth of A. versicolor (Fomicheva et al., 2006). Low pH (below 3.1) prevented growth, while at 25 and 308C growth at pH 10.2 was little different from that at pH 7.4 (Wheeler et al., 1991). Growth of Aspergillus versicolor is greatly affected by modified atmospheres, growth being halved by reduction of O2 concentration to 5%, or by addition of 12–15% CO2, over the range 0.98– 0.90 aw (Magan and Lacey, 1984b). It is a major source of volatile compounds produced by mouldy grain (see Pitt and Hocking, 1997). The decimal reduction time (D10) for Aspergillus versicolor subjected to low dose gamma irradiation was given as 0.45 kGy on grain (O’Neill et al., 1991). Mycotoxins. Aspergillus versicolor is the major producer of sterigmatocystin, a precursor of aflatoxins. Acute oral toxicity is low due to very low solubility in water or gastric juices, so sterigmatocystin is unlikely to be responsible for acute poisoning outbreaks in man or animals (Terao, 1983). Due to this insolubility, experimental doses given to animals are only adsorbed to a small extent. As absorption is dependent on the method of administration, LD50 figures lack accuracy: literature values vary from 60 to 800 mg/kg body weight. However, even low doses can cause tumours in mice (Fujii et al., 1976) and pathological changes to the livers of rats (Terao, 1983). Rats orally administrated sterigmatocystin at the rate of 8.3– 16.6 mg per kg body weight per day for 10 days showed severe fatty acid changes and necrosis of hepatocytes (Aal et al., 1997). Aspergillus versicolor is commonly reported in indoor air, and its presence in damp buildings has been linked to complaints of mouldy odour and eye, nose and throat irritation (Samson et al., 1994). A single dose of 108 A. versicolor spores inhaled by mice induced acute inflammation of the lungs (Jussila et al., 2002). Aerosols of A. versicolor extract administered to mice also caused upper respiratory tract irritation (Korpi et al., 2003). Continuous exposure of baby rats to A. versicolor spores caused pituitary tumors (Sumi et al., 1990, 1994). As a liver carcinogen, sterigmatocystin appears to be only about 1/150th as potent as aflatoxin B1, but this is still much more potent than most other

335

liver carcinogens. Levels as low as 15 mg/day fed continuously, or a single 10 mg dose, caused liver cancer in 30% or more of male Wistar rats (Terao, 1983). Sterigmatocystin also increased the development of intestinal cancer in Mongolian gerbils infected with a known carcinogenic Helicobacter pylori toxin (Ma et al., 2003). Sterigmatocystin has the potential to cause human liver cancer and its occurrence in foods should not be taken lightly. However, unlike aflatoxins and ochratoxin, which occur in the field or during crop drying, sterigmatocystin is only produced in stored grains or cheese, a manufactured product over which control of mould contamination can be achieved (Engel and Teuber, 1980; Veringa et al., 1989). If sterigmatocystin is formed by growth of A. versicolor in rice, most of the toxin is removed at the milling stage (Takahashi et al., 1984). In Canada, the risk of sterigmatocystin formation in stored grains is very low (Mills, 1990). A high proportion of Aspergillus versicolor isolates produce sterigmatocystin, e.g. 18 of 58 tested (Miyaki et al., 1970) or 30 of 32 (Mills and Abramson, 1986). Ecology. A very widely distributed fungus, Aspergillus versicolor, has been reported from most kinds of foods. Although it occurs at harvest in crops such as wheat and barley (see Pitt and Hocking, 1997), it is of much more common occurrence in stored products. Major sources include cereals and oilseeds: wheat, cereal flakes, stored barley, milled rice, oats, rapeseed, sunflower seeds, amaranth seed and vegetable oil (see Pitt and Hocking, 1997). Recent reports include maize (Sepulveda and Piontelli, 2005), flour (Halt et al., 2004), rice (Sakai et al., 2005), soybeans (Sepulveda and Piontelli, 2005) and dried medicinal herbs, especially lemon balm (Janda and Ulfig, 2005). Other sources include peanuts, hazelnuts, walnuts, pistachios, pecans, green coffee beans, frozen meat, biltong peppercorns, spices and health foods (see Pitt and Hocking, 1997). A. versicolor has also been reported in sugar used in the manufacture of soft drinks (Ancasi et al., 2006), fermented and cured meats (Cantoni et al., 2007) and smoked sardines (Nketsia-Tabiri et al., 2003). Aspergillus versicolor has been reported from cheese quite frequently (Minervini et al., 2002 and see also Pitt and Hocking, 1997) and occasionally causes spoilage (Northolt et al., 1980). It is one fungus responsible for decay in fresh breadfruit

336

8 Aspergillus and Related Teleomorphs

(Omobuwajo and Wilcox, 1989) and is also one cause of the ‘‘Rio’’ off-flavour in coffee due to the formation of trichloroanisoles (Liardon et al., 1992). Occurrence of Aspergillus versicolor in Southeast Asian commodities we examined was variable, being quite high in foodstuffs from Indonesia, but low in those from Thailand and the Philippines. This presumably reflected differences in storage times and practice in these countries. Levels in Indonesian pepper (in 12% of all peppercorns examined), kemiri nuts (11%), red beans (10%), cowpeas (5%), coriander and mung beans (2%), peanuts, paddy and milled rice (1%) are probably indicative of growth in storage rather than association with a particular kind of food substrate. In commodities from Thailand, low levels of A. versicolor were found in peanuts, copra, soybeans, paddy and milled rice, wheat and sorghum, while from the Philippines, incidence was limited to low levels in maize, soybeans, mung beans, paddy and milled rice, pepper and sesame seeds (Pitt et al., 1993, 1994, and our unpublished data). References. Raper and Fennell (1965); Domsch et al., (1980); Klich (1993, 2002).

Aspergillus wentii Wehmer

Fig. 8.26

Colonies on CYA 25–35 mm diam, plane or lightly wrinkled, moderately deep to deep, floccose, mycelium white to pale yellow; conidial production moderate, coloured greyish yellow; clear exudate sometimes produced; reverse pale. Colonies on MEA 22–30 mm diam, plane, relatively dense, velutinous; mycelium white; conidial production abundant, orange yellow; reverse pale. Colonies on G25N 30–45 mm diam, plane, deep, with areas of floccose white mycelium, heavily sporing on long stipes; conidia brownish yellow; reverse pale brown. No growth at 5 or 378C. Conidiophores borne from aerial hyphae, on CYA 500–1,200 mm long, on G25N up to 5 mm long, with thin, smooth walls; vesicles nearly spherical, on CYA 25–35 mm diam, on G25N 70–100 mm diam, with metulae and phialides densely packed over the entire surface; metulae 10–18 mm long; phialides ampulliform, 7–12 mm long; conidia spherical to broadly ellipsoidal, 3.5–5.0 mm diam, smooth walled, borne in loose, radiate heads.

Distinctive features. Golden brown colony colour, faster growth as a rule on G25N than CYA and absence of growth at 378C are the principal characters which set Aspergillus wentii apart. Long stipes on G25N, production of metulae and phialides and large smooth-walled conidia are also distinctive. Physiology. Aspergillus wentii is a xerophile which exhibits strong growth in both sugar and salt environments. The optimum aw for growth is near 0.94 in both glucose/fructose and NaCl based media (Andrews and Pitt, 1987). The minimum aw for germination at 258C was reported to be 0.73 aw in glucose/fructose, 0.75 in glycerol and 0.79 aw in NaCl based media (Andrews and Pitt, 1987). A slightly higher limit (0.76 aw in glucose/fructose) was reported by Wheeler et al. (1988b). The minimum aw for germination in glucose/fructose was little affected by temperature: 0.77 at 20 and 308C, 0.79 at 348C and 0.83 at 378C (Wheeler et al., 1988b). Mycotoxins. Emodin is the only mycotoxin of significance produced by Aspergillus wentii (Wells et al., 1975). Administered to day-old cockerels, the oral LD50 was only 3.7 mg/kg, indicating high toxicity (Wells et al., 1975). It has also been reported to be a mutagen (Masuda and Ueno, 1984; Mu¨ller et al., 1996). There are no reports of toxicoses associated with A. wentii, perhaps reflecting the lack of reports of substantial growth, i.e. spoilage, in foods. Ecology. This species has only rarely been reported as a spoilage fungus and is considered to be among the less common major Aspergillus species in foods. Nevertheless, in our experience it is widely distributed. We isolated it in high percentages of many of the Southeast Asian commodities we examined. Twenty three per cent of nearly 500 peanut samples examined contained A. wentii, at up to 100% infection in infected samples and in 6% of all kernels examined. Thirty seven per cent of kemiri nut samples from Indonesia were infected, but the maximum infection level was only 35%, with 6% of nuts infected overall. A wide range of other commodities, maize from all three countries, cashews from Thailand, pepper and soybeans from the Philippines, and mung beans, pepper, coriander and cowpeas from Indonesia, all showed 1–3% overall infection in the particles examined. Low levels of infection were seen in mung beans, soybeans, paddy rice, sorghum, red beans, cassava

8.4

Genus Aspergillus

337

Fig. 8.26 Aspergillus wentii (a) colonies on CYA and MEA, 7 days, 258C; (b, c) heads, bars = 20 mm; (d) conidia, bar = 5 mm

and copra from Thailand and cashews, soybeans, paddy and milled rice from Indonesia (Pitt et al., 1993, 1994, and 1998a). It was also quite common in dried fish (Wheeler et al., 1986). Other records of Aspergillus wentii from foods include peanuts, pistachios, pecans, walnuts, various cereal products, wheat, barley, bread, green

coffee beans, dried beans, ripened raw hams and biltong (see Pitt and Hocking, 1997) and also rice (Taligoola et al., 2004), maize and soybeans (Sepulveda and Piontelli, 2005) and grapes (Pollatro et al., 2005). References. Raper and Fennell (1965); Domsch et al., (1980); Klich (2002).

“This page left intentionally blank.”

Chapter 9

Xerophiles

Xerophilic fungi are distinguished by their ability to grow under conditions of reduced water activity, i.e. to complete their life cycles on substrates that have been dried or concentrated, in the presence of high levels of soluble solids such as salts or sugars. Early usage (Scott, 1957) confined the word ‘‘xerophile’’ to filamentous fungi and used the term ‘‘osmophile’’ for yeasts; the term ‘‘halophile’’ was used rather indiscriminately for moulds, yeasts and bacteria with the ability to grow on concentrated salt solutions. Pitt (1975) discussed the terminology used for fungi which grew at low aw. He pointed out that osmophile was an inappropriate term, because high osmotic pressures were not involved in the growth of these fungi, as they balance the outside environment with internal solutes, maintaining just sufficient osmotic pressure to enable growth. The crucial point was that they preferred to grow at reduced water activities. In the absence of a suitable term for ‘‘lovers of low water activity’’, xerophile seemed the most suitable appellation, for both moulds and yeasts. Some other authors have used the term ‘‘xerotolerant’’ in place of xerophile, arguing that these fungi exhibit ‘‘tolerance of’’ not ‘‘love for’’ reduced aw. However, xerotolerant is an inappropriate term for extremely xerophilic fungi such as Xeromyces bisporus, Chrysosporium fastidium and Basipetospora halophila, which have an absolute requirement for, not tolerance of, reduced aw. Second, although many xerophiles grow optimally at quite high aw, most are basically slowly growing fungi and cannot compete in mixed cultures at high aw levels such as prevail in soils or fresh foods. Survival of xerophiles in nature depends on access to environments of

reduced aw where competition is restricted or eliminated. In this vital sense they are lovers of lowered water activity, i.e. xerophiles. Xerophilic fungi may be defined in a variety of ways, usually relating to minimal or optimal aw for growth. The definition of Pitt (1975) is used here: a xerophile is a fungus capable of growth, under at least one set of conditions, at a water activity below 0.85. This has proved to be a practical working definition. In discussing the use of the word halophile for fungi, Pitt (1975) wrote that it was inappropriate because there were no known fungi with a preference, let alone an obligate requirement, for salt environments. More recently, however, such fungi have been shown unequivocally to exist, exhibiting quite superior growth on media with NaCl as controlling solute. Scopulariopsis halophilica Tubaki was the first; Pitt and Hocking (1985) added Polypaecilum pisce and transferred S. halophilica to Basipetospora (as B. halophila). Perhaps there are others. Such fungi are still classed as xerophiles: if there is a need to differentiate them from other xerophiles, the term ‘‘halophilic xerophile’’ is appropriate (Andrews and Pitt, 1987). In this book, for practical reasons, ‘‘xerophile’’ has a different and much narrower circumscription than the definition given above. Here the definition is derived solely from responses to the standard media. A species has been included, or at least keyed, in this chapter if, after 7 days at 258C, colony diameters on G25N exceed those on CYA and MEA. Many marginally xerophilic fungi which meet the definition of Pitt (1975) do not meet this criterion and have simply been keyed in other appropriate chapters.

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_9, Ó Springer ScienceþBusiness Media, LLC 2009

339

340

9

In this chapter, Eurotium species and a few Aspergilli are keyed out with a miscellaneous group of other fungi. To maintain an orderly presentation, the Eurotium and Aspergillus species have been described in Chapter 8. The miscellaneous fungi are described below, in alphabetical order. Note that in the key which follows, couplets 1–3 are based on growth on G25N, and sort out species which are placed elsewhere, plus one of those

Xerophiles

described below. It is not possible satisfactorily to differentiate the remaining xerophiles on this medium. The subsequent couplets, 4–8, are based on colonies grown on MY50G agar (see Chapter 4). So before entering the key at couplet 4 it will be necessary to inoculate the unknown isolate onto MY50G agar and incubate at 258C for 14 days, and perhaps longer, until fruiting structures differentiate and mature.

Key to xerophilic fungi 1

Bright yellow, barely macroscopic spherical bodies (cleistothecia) visible in the aerial mycelium of colonies on G25N. (Grow on CY20S agar and use key in Chapter 8) Cleistothecia not visible in colonies on G25N

Eurotium 2

2 (1)

Colonies on G25N showing yellow or green conidial colours (Use key in Chapter 8) Colonies on G25N white, brown or black

3 (2)

Colonies on G25N chocolate brown Colonies on G25N white, pale brown or black (Grow on MY50G)

4 (3)

Colonies on MY50G white or pale brown Colonies on MY50G black or with black areas

5 (4)

Colonies on CYA and MEA 10 mm diam or more in 7 days; conidia on MY50G lemon-shaped Colonies on CYA and MEA not exceeding 10 mm diam

Polypaecilum 6

On MY50G, solitary asci produced, containing or releasing mature ascospores in 14 days Mature asci and ascospores not evident in cultures on MY50G in 14 days

Eremascus 7

6 (5)

7 (6)

8 (7)

Intercalary chlamydoconidia and arthroconidia present; aleurioconidia on tiny pedicels or solitary Intercalary chlamydoconidia and arthroconidia absent; aleurioconidia on short conidiophores, sometimes in short chains

Basipetospora was described by Cole and Kendrick (1968) as the anamorph of the Ascomycete genus Monascus (see Chapter 5). The two genera have until recently only been found occurring together, as holomorphic species. Following a light and

Wallemia 4 5 Bettsia (see Chrysosporium)

Colonies on MY50G producing spherical to cylindroidal aleurioconidia or similar conidia in 14 days Colonies on MY50G not producing aleurioconidia in 14 days; 3 celled cleistothecial initials or developing cleistothecia may or may not be evident

9.1 Genus Basipetospora G.T. Cole and W.B. Kendr.

Aspergillus 3

8 Xeromyces

Chrysosporium Basipetospora

scanning electron microscopic study, however, Pitt and Hocking (1985) transferred the salttolerant xerophile Scopulariopsis halophilica Tubaki, a species with no known teleomorph, to Basipetospora. Conidium formation in this fungus appeared to occur by the process characteristic of the Monascus anamorphs. In Basipetospora, conidia (aleurioconidia) are borne in short chains from simple conidiophores,

9.1

Genus Basipetospora

cut off successively from the tip of the conidiophore so that it shortens as the conidia form. Conidia sometimes separate at maturity. The distinction from Chrysosporium is that in the latter genus aleurioconidia are formed solitarily. Moreover conidia borne along the lengths of the vegetative hyphae, i.e. chlamydoconidia and arthroconidia, are not produced by Basipetospora species but are usually present in Chrysosporium species.

Basipetospora halophila (J.F.H. Beyma) Pitt & A.D. Hocking Fig. 9.1 Oospora halophila J.F.H. Beyma Scopulariopsis halophilica Tubaki

No growth on CYA at 5, 25 or 378C or on MEA. Colonies on G25N at 14 days 4–8 mm diam, occasionally 12 mm, of dense and tough mycelium, centrally raised, sulcate or irregularly wrinkled; mycelium persistently white, with little or no sporulation; reverse pale. Colonies on MY50G at 14 days 4–8 mm diam, similar to those on G25N. Colonies on MY5-12 at 14 days 10–16 mm diam, low or umbonate, plane or irregularly wrinkled, of dense mycelium overlaid by floccose to funiculose aerial hyphae; mycelium persistently white, sporulation light to moderate in lower layers of the mycelium; reverse pale to yellow brown. Colonies on MY10-12 at 14 days 18–22 mm diam, similar to those on MY5-12, but growth more rapid and

341

vigorous; sporulation in surface mycelial layers moderate to heavy; mycelium and conidia persistently white; reverse pale to yellow brown. Reproductive structures short, solitary conidiophores borne at irregular intervals along vegetative hyphae, sometimes bearing a short chain of conidia, but more commonly a single developing conidium, shed at maturity and succeeded by another blown out terminally from the conidiophore, the conidiophore shortening a little with each successive conidium; conidiophores often curved, usually cylindrical, 2.0–3.0 mm diam, but sometimes narrowing towards the apex; when young 8–20 mm long, in age down to 3–4 mm long, smooth walled; mature conidia spherical to broadly ellipsoidal or pyriform with a truncate base, 3.5–6.0 mm diam, with heavy walls, smooth to finely roughened, in wet mounts usually solitary, but sometimes in chains of 3 or 4. No teleomorph known. Distinctive features. Although it superficially resembles a Chrysosporium species, Basipetospora halophila is readily distinguished by (1) very slow growth on MY50G; (2) aleurioconidia sometimes in short chains; and (3) the absence of intercalary conidia in the vegetative hyphae. Taxonomy. This distinctive species was described as Oospora halophila by van Beyma (1933). The name was not used by later authors, however. Tubaki (1973) described Scopulariopsis halophilica in terms which did not lead to association with van

Fig. 9.1 Basipetospora halophila (a) colonies on MY5-12, 14 d, 258C; (b,c) conidiophores and aleurioconidia, bars = 20 mm

342

Beyma’s species. Pitt and Hocking (1985) established that O. halophila and S. halophilica were a single species. Studies of S. halophilica by scanning electron microscopy showed that it was not a Scopulariopsis, but belonged in Basipetospora. Physiology. The differences in rates of growth of this fungus on MY50G and MY5-12, media of similar water activity and nutritional status, show clearly that Basipetospora halophila is a true halophilic xerophile. Detailed studies on the water relations of this species confirmed this observation (Andrews and Pitt, 1987; Wheeler et al., 1988c). Three isolates of B. halophila germinated in 6–11 days on a saturated NaCl medium (0.747 aw). Germination was slower on media containing glucose/fructose or glycerol as controlling solute; however, one isolate germinated at 0.73 aw in 40 days on glucose/fructose. The lowest aw for germination on glycerol media was 0.78. Growth rates of three isolates were similar, but showed marked differences on the different media: the radial growth rate on NaCl at 0.75 aw (10 mm/h) was comparable with the maximum rate observed on glucose/fructose at 0.90 aw. Maximum growth rates on NaCl, ca. 30 mm/h, occurred at 0.87–0.88 aw. Strong growth of Basipetospora on NaCl based media was also demonstrated by Diyaolu and Adebajo (1994). The optimum temperature for growth of Basipetospora halophila is near 308C, although it was more tolerant of high aw conditions at lower temperatures, growing at 0.99 aw on NaCl based media at 258C. At 378C, B. halophila demonstrated slow growth on NaCl based media, and over only a limited aw range, 0.86–0.77 aw (Wheeler et al., 1988c). The maximum temperature for growth is probably not much higher than 378C. The minimum temperature for growth is not known. Basipetospora halophila is well adapted to its ecological niche – dried, salted seafoods and similar substrates. Studies on competition between common xerophiles such as Eurotium species and halophilic xerophiles growing together on salted fish have clearly shown that B. halophila and the other halophilic xerophile, Polypaecilum pisce, can outgrow the Eurotia below about 0.87 aw (Wheeler and Hocking, 1993). Mycotoxins. No mycotoxins are known from this species.

9

Xerophiles

Ecology. Basipetospora halophila is a halophilic xerophile which is restricted to salty environments. Most isolates of B. halophila have come from salted, dried fish: it has been reported from dried or cured fish from Japan, Indonesia, the Philippines, Sri Lanka and Nigeria (see Pitt and Hocking, 1997). Other sources include dried food grade seaweed in Japan, a gelatine hydrolysate (Tubaki, 1973) and sea salt (as Scopulariopsis halophilica; CBS, 2007). Reference. Pitt and Hocking (1985).

9.2 Genus Chrysosporium Corda Chrysosporium is a genus of Hyphomycetes characterised by the formation of solitary, hyaline, smooth walled aleurioconidia. These are produced on the sides of vegetative hyphae either directly (sessile) or on small pedicels. Some species also produce similar conidia terminally on hyphae (terminal chlamydoconidia). In a few species, the vegetative hyphae themselves also differentiate either partially or almost totally into conidia: some are nearly spherical (intercalary chlamydoconidia); others are produced from unswollen hyphal segments (arthroconidia). These two spore types may intergrade. Speciation in this genus largely depends on the size, shape and proportion of these various types of conidia. Most Chrysosporium species are found in soil, or on the hair or skin of animals, and are dependent on high water activities for growth. Some are pathogenic. Species which occur in foods, however, are not pathogenic and are xerophiles. However, molecular work in our laboratory has indicated that they are genetically related to some of the nonxerophilic Chrysosporium species (N. Tran-Dinh, unpublished). Most Chrysosporium species are strictly anamorphic. Some of the dermatophyte species form teleomorphs in Gymnoascus, Arthroderma and other related genera, outside the scope of this work. One of the foodborne species, C. farinicola, sometimes produces a teleomorph classified in Bettsia Skou, an Ascomycete which forms unique black cleistothecia. Molecular studies have shown that it belongs to the Eurotiales (N. Tran-Dinh, unpublished). Skou (1992)

9.2

Genus Chrysosporium

343

described seven new species and three new varieties of Chrysosporium from the pollen and nectar of mason bees. Although not all of Skou’s species are molecularly distinct (N. Tran-Dinh, unpublished), his work indicated that the natural habitat for xerophilic Chrysosporium species is associated with bees, beehives and honey. Kinderlerer (1995) developed a medium, Czapek casein 50% glucose agar (CZC50G) to

assist in the identification of xerophilic foodborne Chrysosporium species. The key to xerophilic Chrysosporium species deals only with the four species most commonly encountered in foods and is based on growth on MY50G agar for 7 days at 258C. Confirmation of identity may depend on colony maturation which can take 2–3 weeks.

Key to xerophilic Chrysosporium species 1

2 (1)

Colonies on MY50G exceeding 15 mm diam in 7 days; predominant conidial type solitary aleurioconidia Colonies on MY50G not exceeding 15 mm diam in 7 days, and not usually exceeding 30 mm in 14 days; predominant conidial type intercalary chlamydoconidia and arthroconidia

Colonies on MY50G pale yellow or brown with a yellow brown reverse Colonies on MY50G persistently white with a pale reverse, or showing dark grey to black areas

Chrysosporium farinicola (Burnside) Skou Fig. 9.2 Ovularia farinicola Burnside

Teleomorph: Bettsia alvei (Betts) Skou Pericystis alvei Betts

No growth on CYA at 5, 25 or 378C or on MEA. Colonies on G25N 10–20 mm diam, of low, white mycelium, or if the teleomorph is present, centrally grey from the production of immature cleistothecia; margins fimbriate; reverse pale, but grey centrally if the Bettsia state is being produced. Colonies on MY50G growing relatively rapidly, at 7 days 20–30 mm diam, low, plane, persistently white, or showing sectors becoming translucent or greyish if the Bettsia state is present, reverse pale beneath white areas, but darker beneath grey sectors; at 14 days, 40–65 mm diam, colonies remaining white if only the anamorph is present, but with dark grey to black areas in the presence of the teleomorph; reverse pale, or dark under the teleomorph (Fig. 9.2). Reproduction on MY50G predominantly solitary aleurioconidia, broadly ellipsoidal to pyriform, thick walled, highly refractile, 7–11 mm long. Fertile hyphae mostly undifferentiated, dissolving in age. In isolates producing the teleomorph, areas or sectors of translucent growth developing. At maturity such areas becoming grey and then black as small

2 C. inops C. xerophilum C. fastidium C. farinicola

cleistothecia form. Cleistothecia formed on MY50G at 258C, dark brown to black, usually maturing only after several weeks at 15–258C, 25–60 mm diam, with walls thin and smooth, and without internal structure; initials a row of three short cells, 12–18  6–8 mm overall, adhering to the cleistothecial wall as a distinctive appendage; ascospores not liberated readily, spherical, 5–6 mm diam, with dark walls, smooth to minutely roughened. Distinctive features. Chrysosporium farinicola is distinguished from C. fastidium by the following features: (1) colonies on MY50G grow more rapidly; (2) colonies on G25N and MY50G remain pure white, with reverses virtually uncoloured; and (3) terminal chlamydoconidia are often larger, up to 13–18 mm in diameter. In addition, some isolates of C. farinicola produce the Bettsia state. No teleomorph is known for C. fastidium. Mycotoxins. No mycotoxins are known from this species. Physiology. No studies on the water relations of this species are known to us, but Chrysosporium farinicola would be expected to resemble C. fastidium in its response to aw. Studies on recovery of heat stressed C. farinicola aleurioconidia showed that older cultures were less heat sensitive than 14 day cultures (Beuchat and Pitt, 1990b). There was less than one log reduction in 20 day old aleurioconidia heated at 568C for 10 min. Aleurioconidia survived

344

9

Xerophiles

Fig. 9.2 Chrysosporium farinicola and Bettsia alvei (a) colonies of the Chrysosporium state alone and (b) with the Bettsia state also present, both on MY50G, 14 d, 258C; (c) aleurioconidia, bar = 10 mm; (d) mature cleistothecia-containing ascospores, bar = 25 mm; (e) ascospores, bar = 5 mm

well in maize starch at 0.43 and 0.71 aw at 1 and 258C, showing no reduction in numbers after 45 days storage. Heat stressed C. farinicola aleurioconidia grew better on glucose based media than on media containing sorbitol, glycerol or NaCl (Beuchat and Pitt, 1990b). Isolates of C. farinicola were able to grow in the presence of 1.05–1.76 % (w/v) ethanol and at temperatures ranging from 4 to 258C (Kinderlerer, 1997). Growth in controlled atmosphere was recorded in a mixture of 10% CO2 and 20% O2 or 6% O2 (Kinderlerer, 1997). Weak protease activity and moderate lipolytic activity were reported for C. farinicola from Kenyan coconut (Ismail, 2001). Ecology. Chrysosporium farinicola has been isolated from a range of low aw substrates, including

honeycomb, prunes and prune processing equipment, sultanas, mixed dried fruit, chocolate, jelly crystals and coconut from Australia, the United Kingdom, Sri Lanka, Czechoslovakia, Denmark and Kenya (Kinderlerer, 1997; Ismail, 2001; Food Science Australia, 2006; CBS, 2007). In coconut, C. farinicola can cause cheesy butyric spoilage (Kinderlerer, 1984a). References. Skou (1975); van Oorschot (1980).

Chrysosporium fastidium Pitt

Fig. 9.3

No growth on CYA at 5, 25 or 378C, or on MEA. Colonies on G25N 1–5 mm diam, of low, dense,

9.2

Genus Chrysosporium

345

Fig. 9.3 Chrysosporium fastidium (a) colonies on MY50G, 14 d, 258C; (b,c) aleurioconidia, borne on pedicels, bar = 10 mm

white mycelium. Colonies on MY50G at 7 days 15– 22 mm diam, low, plane and sparse, pale yellow or brown, reverse yellow brown; at 14 days, 35–42 mm diam, low and plane, margins sparse and fimbriate, white, centres more dense, dull yellow; reverse yellow to pale brown. Reproduction on MY50G predominantly by smooth walled aleurioconidia borne singly on short pedicels or less commonly sessile, spheroidal (oblate or prolate) to broadly ellipsoidal, 6–9  5–8 mm, in age released by dissolution of the pedicels; terminal chlamydoconidia, spherical to pyriform or pedunculate, 8–12  6–10 mm, also produced, but intercalary chlamydoconidia and arthroconidia rare. No teleomorph known. Distinctive features. Chrysosporium fastidium forms dull yellow to yellow brown colonies; conidia are predominantly aleurioconidia with few intercalary chlamydoconidia or arthroconidia produced on G25N or MY50G. It most closely resembles C. farinicola from which is can be distinguished by colony colour (C. farinicola is persistently white or has dark grey areas where the Bettsia teleomorph is formed), and slower growth on MY50G and Czapek casein 50% glucose agar (Kinderlerer, 1995). Taxonomy. Van Oorschot (1980) placed Chrysosporium fastidium in synonymy with C. farinicola, but this is not accepted here. The two species are readily distinguished and only C. farinicola forms a teleomorph.

Physiology. A mesophilic obligate xerophile, Chrysosporium fastidium has a maximum aw for growth of 0.98 and a minimum of 0.69 (Pitt and Christian, 1968). It does not utilise nitrate and appears to require accessory factors for growth. Conidia have an unexceptional heat resistance (Pitt and Christian, 1970). Mycotoxins. No mycotoxins are known from this species. Ecology. This species has been repeatedly isolated from prunes (dried and high moisture) and prune-processing machinery in N.S.W., Australia. There are no records of isolation from other substrates or other locations, indicating that Chrysosporium fastidium is a rare species with a restricted habitat. Reference. Pitt (1966).

Chrysosporium inops J.W. Carmich.

Fig. 9.4

Colonies on CYA and MEA microscopic or up to 3 mm diam. Colonies on G25N 2–9 mm diam, varying from low and translucent to deep and floccose, white; reverse pale to amber or duller yellow brown. No growth on CYA at 5 or 378C. Colonies on MY50G at 7 days 6–10 mm diam, at 14 days 12–20 mm diam, varying from low, sparse and translucent to moderately deep, dense and with a floccose surface, white or if translucent, uncoloured; reverse uncoloured to pale yellow brown.

346

9

Xerophiles

Fig. 9.4 Chrysosporium inops (a) colonies on MY50G, 14 d, 258C; (b,c,d) chains of arthroconidia, borne by hyphal transformation, bars = 10 mm

Reproductive structures on G25N or MY50G at 7 days primarily short chains of chlamydoconidia and arthroconidia, borne by retrogressive differentiation from hyphal tips and as intercalary chains; some terminal chlamydoconidia also present; at maturity on MY50G (2–4 weeks), clusters of chlamydoconidia and arthroconidia also present, formed by retrogressive differentiation of groups of short branching hyphae with a lateral stipe as their common origin. Chlamydoconidia spherical, 4–7(–10) mm diam; arthroconidia cylindroidal or doliiform (barrel shaped), 3–8  3–6 mm, those of greater width intergrading with chlamydoconidia; aleurioconidia, the conidial type characteristic of Chrysosporium, uncommon, ellipsoidal to pyriform, 5–8 mm diam or longer. Large chlamydoconidia, up to 25 mm diam, with walls up to 2 mm thick, produced by some isolates. Teleomorph unknown. Distinctive features. In culture, Chrysosporium inops (and C. xerophilum; see below) differ from C. fastidium and C. farinicola by slower growth rates, especially on MY50G. The two species under consideration here also produce predominantly chlamydoconidia and arthroconidia. On CZC50G medium, some isolates of C. inops produce a red reverse, but most are yellow (Kinderlerer, 1995). C. inops grows more slowly than C. xerophilum on this medium. In young colonies (7 days),

terminal chlamydoconidia are often the dominant conidium type in C. inops. Aleurioconidia are rare. Physiology. Van Oorschot (1980) reported that Chrysosporium inops had a minimum growth temperature of 208C, an optimum of 258C and a maximum of 308C. However, her data were obtained on media of very high aw, and limits are undoubtedly wider under optimal conditions. The water relations of this species have not been studied in detail, but isolations in our laboratory have come from growth under controlled aw levels as low as 0.72. It is clear, contrary to previous reports (Pitt and Christian, 1968; van Oorschot, 1980), that C. inops is an extreme xerophile. However, it is also able to grow (very slowly) at the very high aw of CYA and MEA. Arthroconidia of Chrysosporium inops are moderately heat resistant, and their heat resistance increases with maturity. The decimal reduction time at 668C (D66) was 1.67 min for 3 week old spores, 1.95 min for 4 week old spores and 5.49 min for 6 week old spores (Kinderlerer, 1996). A small population of heat resistant arthroconidia survived heating at 668C for 1 h, representing approximately 0.001% of the population (Kinderlerer, 1996). The heating menstruum in these experiments was 0.1% peptone. A higher heat resistance could be expected in high sugar environments. Chrysosporium inops can produce ethanol from glucose (Kinderlerer, 1987). Its ability to utilise the

9.3

Genus Eremascus

Embden–Meyerhof–Parnas pathway may explain why this species and the closely related C. xerophilum may become dominant in low aw foods stored at low oxygen tension (Kinderlerer, 1987). Mycotoxins. No mycotoxins are known from this species. Ecology. This appears to be an uncommon species, but as with other xerophilic Chrysosporia, this may only reflect the need for well-chosen isolation techniques and identification procedures. We have isolated Chrysosporium inops from a variety of spoiled products: nutmeg powder, Chinese five spice powder, mixed spice powder, chopped Chinese dates and a gelatine confection made in a starch mould. In the latter case, 30 tonnes of starch were heavily contaminated with this fungus, causing a serious loss of confectionery products. There was no evidence that the starch had ever been more than marginally above a safe aw level. C. inops has also been isolated from table jelly (Kinderlerer, 1987) and spoiled chocolate and hazelnuts in the United Kingdom (J.L. Kinderlerer, unpublished). Christensen (1978b) reported C. inops from samples of maize stored for 12 months or more at low moisture in tightly closed containers. His identification was in error (see below); however, we have recovered C. inops from a sample of safflower seeds stored in a similar way in our laboratory. Additional species. Chrysosporium xerophilum Pitt is closely related to C. inops. It differs by (1) faster growth, especially on MY50G, where colonies at 7 days are 10–15 mm diam and at 14 days 25–32 mm diam; (2) higher numbers of aleurioconidia, which measure 7–8  5–7 mm; (3) larger terminal chlamydoconidia, 10–12(–15) mm diam; and (4) at maturity, almost complete differentiation of vegetative hyphae into intercalary chlamydoconidia and arthroconidia, even aleurioconidium pedicels often becoming thick walled spores. CZC50G medium may be useful in differentiating these two species (Kinderlerer, 1995). A mesophilic xerophile, Chrysosporium xerophilum has a minimum aw for growth of 0.71 (Pitt and Christian, 1968) and, like C. inops, a high maximum limit. Although most conidia of this species have a low heat resistance, surviving less than 10 min at 608C, a small proportion of conidia appear to be quite resistant, surviving at least 708C for 10 min (Pitt and Christian, 1970). A similar

347

effect was observed for conidia of C. inops (Kinderlerer, 1996). For a long time this species was known only from the type isolate, from moist Australian prunes. Christensen (1978b) isolated it (reported as Chrysosporium inops) from US maize and other oilseed samples which had been stored at moisture contents of 15–16% in sealed containers for periods of 1–10 years. Other isolates have come from coconut and chocolate (Kinderlerer, 1997) in the United Kingdom and Australia (our laboratory). In spoiled coconut, C. xerophilum formed a wide range of volatile compounds including aliphatic methyl ketones, esters and secondary alcohols as well as free medium chain length fatty acids (Kinderlerer et al., 1988). References. Pitt (1966); van Oorschot (1980).

9.3 Genus Eremascus Eidam Eremascus is distinguished by the formation of asci which are borne singly from undifferentiated hyphae without any surrounding wall or hyphal network. In this characteristic, it resembles Byssochlamys. However, no anamorph is produced as a rule. Molecular analysis of the ITS region places Eremascus in the Eurotiomycetidae along with Aspergillus, Penicillium and other genera described here (Berbee and Taylor, 1992). Eremascus species are strict xerophiles, growing only at reduced aw. Colonies are floccose and remain persistently white, and apparently sterile. Under the compound microscope, abundant asci can be seen. There are two closely related species, E. albus and E. fertilis, distinguished by differences in the cells supporting the asci. B